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LACCASE IN ORGANIC SYNTHESIS AND ITS APPLICATIONS
A Dissertation Presented to
The Academic Faculty
by
Suteera Witayakran
In Partial Fulfillment of the Requirements for the Degree
Doctor of Philosophy in the School of Chemistry and Biochemistry
Georgia Institute of Technology December 2008
LACCASE IN ORGANIC SYNTHESIS AND ITS APPLICATIONS
Approved by: Dr. Arthur J. Ragauskas, Advisor School of Chemistry and Biochemistry Georgia Institute of Technology
Dr. John Cairney School of Biology Georgia Institute of Technology
Dr. David M. Collard School of Chemistry and Biochemistry Georgia Institute of Technology
Dr. Preet M. Singh School of Mechanical Engineering Georgia Institute of Technology
Dr. Uwe H. F. Bunz School of Chemistry and Biochemistry Georgia Institute of Technology
Date Approved: October 22, 2008
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ACKNOWLEDGEMENTS
First, I would like to thank my advisor, Dr. Arthur J. Ragauskas, for his
instruction, encouragement, advice, and support throughout my graduate education. I also
would like to thank my thesis committee, Dr. David M. Collard, Dr. Uwe H. F. Bunz, Dr.
John Cairney, and Dr. Preet M. Singh for their insightful comments and support
throughout this project.
I would like to especially thank Dr. Leslie Gelbaum. His guidance and support in
teaching me the techniques of NMR spectroscopy has helped me immensely in this
research. I also would like to thank Dr. Robert Braga for conducting the IR experiments
in the synthesis of p-naphthoquinone project.
I also appreciate the assistance of all of my co-workers in Dr. Ragauskas’ lab that
allowed me to complete my research tasks, and especially, I would like to thank the
following colleagues:
Dr. Yunqiao Pu for his kind help in NMR spectroscopy; Lenong Allison for her
strong support through my study; Dr. Kristina Knutson for her training in how to measure
laccase activity; Dong Ho Kim for his assistance in bulk acid group testing and handsheet
making; Shaobo Pan and Shoujian Hu for their kind help in taking SEM images; Dr. Nan
Jiang for his helpful advice for my synthetic work; Lee Goetz and Poulomi Sannigrahi for
their friendship and support during my study; Rajalaxmi Dash for her help in FT-IR
experiments; and Abdullah Zettili, summer undergrad student, for helping me in the
synthesis of ortho-naphthoquinone project.
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I am pleased to acknowledge the IPST endowment fund and the Royal Thai
Government Scholarship for providing me with financial support throughout my years of
study. Ms. Tuwanda Strowbridge is appreciated for her guidance and help in the paper
testing lab.
Lastly, I would like to thank my parents, family, and friends for always
supporting and encouraging me in my education.
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TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS……………………………………………………………..iii
LIST OF TABLES…………………………………………………………………….. xii
LIST OF FIGURES………………………………………………………………….…xiv
LIST OF EQUATIONS………………………………………………………………..xxii
NOMENCLATURE…………………………………………………………………. xxiii
SUMMARY…………………………………………………………………………...xxvi
CHAPTER
1 INTRODUCTION………………………………………………………………. 1
1.1 Introduction……………………………………………………………… 1
1.2 Objectives………………………………………………………………...3
2 LITERATURE REVIEW………………………………………………………...5
2.1 Green Chemistry………………………………………………………… 5
2.1.1 Definition of Green Chemistry……………………………………. 5
2.1.2 Twelve Principles of Green Chemistry……………………………. 5
2.2 Water as Solvent in Organic Synthesis………………………………….. 8
2.2.1 Diels-Alder Reaction ……………………………………………..10
2.2.1.1 Quinone Diels-Alder reaction……………………………… 15
2.2.1.2 Uncatalyzed Diels-Alder Reaction in Aqueous Medium….. 22
2.2.1.3 Lewis-Acid-Catalyzed Diels-Alder Reaction in
Aqueous Medium…………………………………………. 30
2.3 Biocatalysis…………………………………………………………….. 35
2.3.1 Enzymes………………………………………………………….. 35
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2.3.1.1 Nomenclature and Classification…………………………... 35
2.3.1.2 Enzyme Mechanism………………………………………... 36
2.3.1.3 Enzyme Kinetics…………………………………………… 38
2.3.1.4 Advantages and Disadvantages of Biocatalyst…………….. 40
2.3.2 Enzymes in Domino Reactions…………………………………... 41
2.3.2.1 Enzyme-Triggered Diels-Alder Reaction………………….. 42
2.3.2.2 Enzyme-Triggered Rearrangement………………………… 44
2.3.2.3 Enzyme-Triggered Fragmentation…………………………. 47
2.3.2.4 Enzyme-Triggered Intramolecular Substitution Affecting
Cyclization…………………………………………………. 48
2.3.2.5 Enzyme-Triggered Other Type of Ractions………………... 52
2.3.2.6 Multienzymatic One Pot Reaction…………………………. 53
2.4 Laccase…………………………………………………………………. 56
2.4.1 Distribution in Nature……………………………………………. 56
2.4.2 Laccase Structure………………………………………………… 57
2.4.3 Catalytic Mechanism and Properties……………………………...59
2.4.4 Laccase in Organic Synthesis……………………………………. 64
2.4.4.1 Laccase-Catalyzed Oxidation Reaction……………………. 65
2.4.4.2 Laccase-Mediated Formation of Intermediate Quinones
In Organic Synthesis……………………………………….. 80
2.4.4.3 Laccase-Catalyzed Polymerization Reaction………………. 84
2.4.5 Laccase in Fiber Modification…………………………………… 88
2.4.5.1 Lignocellulosic Fibers……………………………………… 89
2.4.5.2 Laccase Application in Fiber Modification………………. 105
2.4.6 Conclusions……………………………………………………... 114
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2.5 Lipase…………………………………………………………………. 115
2.5.1 A General Account………………………………………………115
2.5.2 Lipase-Catalyzed Michael Reaction……………………………. 119
3 EXPERIMENTAL MATERIALS AND PROCEDURES…………………….124
3.1 Materials……………………………………………………………… 124
3.1.1 Chemicals……………………………………………………….. 124
3.1.2 Enzymes………………………………………………………… 124
3.1.3 Pulp……………………………………………………………... 126
3.2 Experimantal Procedures for the Use of Laccase in Organic
Synthesis……………………………………………………………… 127
3.2.1 General Information…………………………………………….. 127
3.2.2 Analytical Analysis Procedures………………………………… 128
3.2.3 General Procedure of the Synthesis of 1,4-Naphthoquinones
and Related Structures…………………………………………...129
3.2.4 General Procedure of the Synthesis of o-Naphthoquinones……..130
3.2.5 General Procedure of the Synthesis of Benzofuran Derivatives
via Laccase Oxidation-Michael Addition………………………. 131
3.2.6 General Procedure of the Synthesis of Benzofuran Derivatives
Using Laccase-Lipase Co-catalytic System…………………….. 132
3.2.7 General Procedure of the Reaction of Catechols and Anilines
Catalyzed by Laccase-Lipase Co-catalytic System……………...132
3.3 Experimental Procedures for the Use of Laccase in Fiber
Modification…………………………………………………………...133
3.3.1 Pulp Treatment………………………………………………….. 133
3.3.2 Bulk Acid Group Measurement………………………………… 133
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3.3.3 Pulp Refining and Handsheet Formation……………………….. 135
3.3.4 Paper Physical Tests……………………………………………..137
3.3.5 Nitrogen Analysis………………………………………………..138
3.3.6 Scanning Electron Microscope (SEM)…………………………..139
4 ONE POT SYNTHESIS OF 1,4-NAPHTHOQUINONES AND RELATED
STRUCTURES WITH LACCASE…………………………………………... 140
4.1 Introduction…………………………………………………………… 140
4.2 Experimental Section…………………………………………………. 143
4.2.1 Meterials…………………………………………………………143
4.2.2 Enzyme Assay…………………………………………………... 144
4.2.3 General Procedure for the Study of the Effect of Laccase Dose
and Temperature………………………………………………... 144
4.2.4 General Procedure of the Synthesis of 1,4-Naphthoquinones
and Related Structures…………………………………………...145
4.2.5 Product Characterization………………………………………... 146
4.3 Results and Discussion………………………………………………...152
4.3.1 Preliminary Study of the Reaction System……………………... 152
4.3.2 The Effect of Laccase Dose…………………………………….. 154
4.3.3 The Effect of Temperature……………………………………… 157
4.3.4 The Reaction of p-Hydroquinone and Dienes…………………...159
4.4 Conclusions…………………………………………………………… 162
5 LACCASE-GENERATED QUINONES IN 1,2-NAPHTHOQUINONE
SYNTHESIS VIA DIELS-ALDER REACTION............................................... 163
5.1 Introduction…………………………………………………………… 163
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5.2 Experimental Section…………………………………………………. 166
5.2.1 Enzyme Assay…………………………………………………... 166
5.2.2 General Procedure of the Synthesis of o-Naphthoquinones……..166
5.2.3 Typical Experimental Procedure for p-Naphthoquinone
Synthesis………………………………………………………... 166
5.2.4 Product Characterization………………………………………... 167
5.3 Results and Discussion………………………………………………...171
5.4 Conclusions…………………………………………………………… 180
6 CASCADE SYNTHESIS OF BENZOFURAN DERIVATIVES VIA
LACCASE OXIDATION-MICHAEL ADDITION………………………….. 181
6.1 Introduction…………………………………………………………… 181
6.2 Experimental Section…………………………………………………. 183
6.2.1 General Information…………………………………………….. 183
6.2.2 Enzyme Assay…………………………………………………... 183
6.2.3 General Procedure of the Synthesis of Benzofuran Derivatives
via Laccase Oxidation-Michael Addition………………………. 183
6.2.4 Product Characterization………………………………………... 184
6.3 Results and Discussion………………………………………………...186
6.3.1 Preliminary Study and the Effect of pH on the Reaction System.186
6.3.2 The Effect of the Lewis Bases on the Reaction System…………187
6.3.3 The Effect of the Lewis Acids on the Reaction System…………189
6.3.4 The Synthesis of Benzofuran Derivatives………………………. 190
6.3.5 The Recyclability of the Laccase/Sc(OTf)3-Catalytic System….. 194
6.4 Conclusions…………………………………………………………… 196
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7 CO-CATALYTIC ENZYME SYSTEM FOR THE MICHAEL ADDITION
REACTION OF IN SITU-GENERATED ORTHO QUINONES……………. 197
7.1 Introduction…………………………………………………………… 197
7.2 Experimental Section…………………………………………………. 199
7.2.1 General Information…………………………………………….. 199
7.2.2 Enzyme Assay…………………………………………………... 200
7.2.3 General Procedure of the Synthesis of Benzofuran Derivatives
Using Laccase-Lipase Co-catalytic System…………………….. 200
7.2.4 Procedure for the Study of the Reaction of 5a and 8a
(with and without Lipase)………………………………………. 200
7.2.5 General Procedure of the Reaction of Catechols and Anilines
Catalyzed by Laccase-Lipase Co-catalytic System……………...201
7.2.6 Product Characterization………………………………………... 202
7.3 Results and Discussion……………………………………………….. 205
7.3.1 Laccase-Lipase Co-Catalytic System for the Reaction of
Catechols and 1,3-Dicarbonyl Compounds……………………...205
7.3.2 Laccase-Lipase Co-Catalytic System for the Reaction of
Catechols and Anilines…………………………………………. 213
7.4 Conclusions…………………………………………………………… 217
8 MODIFICATION OF HIGH-LIGNIN CONTENT SOFTWOOD KRAFT
PULP WITH LACCASE AND AMINO ACIDS……………………………. 218
8.1 Introduction…………………………………………………………… 218
8.2 Experimental Section…………………………………………………. 221
8.2.1 Materials…………………………………………………………221
8.2.2 Enzyme Assay…………………………………………………... 222
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8.2.3 Pulp Treatment………………………………………………….. 222
8.2.4 Bulk Acid Group Measurement………………………………… 222
8.2.5 Paper Testing…………………………………………………….223
8.3 Results and Discussion………………………………………………...223
8.3.1 Preliminary Study of the Grafting Condition…………………… 223
8.3.2 The Effect of Amino Acid on the Modifying Fibers…………….226
8.3.3 The Effect of Laccase Dose…………………………………….. 230
8.3.4 Nitrogen Content of Laccase-His Treated Pulp………………… 231
8.3.5 Paper Strength Properties……………………………………….. 232
8.4 Conclusions…………………………………………………………… 236
9 OVERALL CONCLUSIONS………………………………………………… 237
10 RECOMMENDATIONS FOR FUTURE WORK…………………………….241
APPENDIX A: NMR AND IR SPECTRA OF NEW COMPOUNDS……………...242
APPENDIX B: COPYRIGHT PERMISSION……………………………………… 273
APPENDIX C: TENSILE AND TEAR STRENGTH……………………………… 280
REFERENCES……………………………………………………………………….. 286
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LIST OF TABLES
Page
Table 1. The examples of uncatalyzed and catalyzed quinone Diels-Alder reaction. ...... 17 Table 2. Example of the hetero Diels-Alder reactions studied by Lubineau et al. ........... 29 Table 3. Some examples of laccase mediated transformation of natural compounds. ..... 70 Table 4. Substrates, reaction conditions, and products from laccase catalyzed polymerization reactions................................................................................................... 85 Table 5. The degree of polymerization and percentage of the major hemicelluloses in softwoods and hardwoods................................................................................................. 92 Table 6. The percentage of different lignin linkages in hardwood and softwood.[279,284]........................................................................................................................................... 96 Table 7. Yield values for individual pulp components after kraft pulping of Scots pine (a softwood) and birch (a hardwood).................................................................................. 103 Table 8. Paper strength test result for high lignin kraft pulp treated with laccase and phenolic acids.................................................................................................................. 110 Table 9. The reaction of p-hydroquinones and dienes.................................................... 161 Table 10. Preliminary study of the laccase-catalyzed reaction of catechol (5a) and 2,3-dimethyl-1,3-butadiene (2a) in aqueous medium ........................................................... 171 Table 11. Solvent effect on the laccase-catalyzed reaction of catechol (5a) and 2,3-dimethyl-1,3-butadiene (2a)............................................................................................ 173 Table 12. The study of laccase-catalyzed reaction of 2a with a variety of catechol substrates in aqueous medium ........................................................................................ 175 Table 13. The study of laccase-catalyzed reaction of 4-methylcatechol with a variety of dienes in aqueous medium.............................................................................................. 177 Table 14. The study of laccase-catalyzed reaction of 1-acetoxy-1,3-butadiene with a variety of 1,4-benzohydroquinone in aqueous medium at 55 oC.................................... 179 Table 15. The effect of pH on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a) ........................................................................................................... 187
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Table 16. The effect of Lewis bases on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a) ................................................................... 188 Table 17. The effect of Lewis acids on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a)............................................................................................. 190 Table 18. The study of the laccase/Sc(OTf)3-catalyzed reaction of catechols and 1,3-dicarbonyl compounds for benzofuran synthesis............................................................ 192 Table 19. 1H and 13C assignment and HMBC correlations for compound 9da ............... 194 Table 20. Recycling of the laccase/Sc(OTf)3 catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c)........................................................ 195 Table 21. 1H and 13C assignments and HMBC correlation for compound 11b, 11c, and 11d................................................................................................................................... 204 Table 22. Reaction of catechol (5a) and acetylacetone (8a) in the presence of laccase with a variety of lipases. ......................................................................................................... 207 Table 23. The study of the laccase/lipase catalyzed reaction of catechols and 1,3-dicarbonyl compounds in aqueous medium.................................................................... 211 Table 24. Recycling of the laccase/lipase co-catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c)........................................................ 212 Table 25. Reactions of catechol and anilines in the presence of laccase,with (or without) lipase PS in aqueous medium. ........................................................................................ 216
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LIST OF FIGURES
Page
Figure 1. Diels-Alder reaction of 1,3-butadiene with ethylene. ....................................... 10 Figure 2. Schematic drawing of the molecular orbitals of alkenes and conjugated dienes and the orbital interaction for normal and inverse electron demand Diels-Alder reactions. ........................................................................................................................... 12 Figure 3. Example of the regioselectivity of normal electron-demand Diels-Alder reaction controlled by the orbital coefficients of the atoms forming the σ-bonds.......................... 13 Figure 4. The endo and exo approach of the Diels-Alder reaction between piperylene and acrolein and the secondary orbital interaction in the endo transition state. ...................... 14 Figure 5. The quinone Diels-Alder (QDA) reaction......................................................... 15 Figure 6. A quinone-based Diels-Alder reaction as the key step in the total synthesis of the steroid hormones cortisone and cholesterol. ............................................................... 16 Figure 7. A Diels-Alder reaction of quinone and a vinyl cyclohexene as the key step in the total synthesis of forskolin derivative. ........................................................................ 20 Figure 8. A Diels-Alder reaction of Danishefsky-type diene and quinone in the presence of the Mikami’s catalyst for the total synthesis of (-)-colombiasin A. ............................. 21 Figure 9. A Diels-Alder reaction of 1,3-diene and 1,4-benzoquinone in the presence of the Mikami’s catalyst as a key step for the total synthesis of ibogamine. ........................ 21 Figure 10. Cr-catalyzed asymmetric quinone Diels-Alder reaction as a key step for the total syntheses of (-)-colombiasin A and (-)-Elisapterosin B. .......................................... 22 Figure 11. Diels-Alder reaction between cyclopentadiene and methyl vinyl ketone in water and organic solvents................................................................................................ 23 Figure 12. Relative reaction rate (kwater/ kn-hexane) of Diels-Alder reaction between 2,3-dimethyl-1,3-butadiene and N-alkylmaleimides.............................................................. 24 Figure 13. Diels-Alder reaction between trans,trans-2,4-hexadienyl acetate and N-propylmaleimide under various conditions....................................................................... 25 Figure 14. Diels-Alder reaction between α,β-unsaturated ketoaldehyde and ethyl 4-methyl-3,5-hexadienoate................................................................................................... 26
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Figure 15. Intramolecular hetero Diles-Alder reaction of N-acylnitroso compound........ 27 Figure 16. Intramolecular imino-Diels-Alder reactions.................................................... 30 Figure 17. Complexation of Cu(L-abrine) catalyst and 3-phenyl-1-(2-pyridyl)-2-propen-1-one ................................................................................................................................. 31 Figure 18. The enantioselectivity of copper (L-arabine) catalyzed Diels-Alder reactions of 3-phenyl-1-(2-pyridyl)-2-propen-1-one with cyclopentadiene .................................... 31 Figure 19. The aqueous aza-Diels-Alder reaction using lanthanide triflate ..................... 32 Figure 20. Yb(OTf)3-catalyzed Diels-Alder reaction between N-benzylideneaniline as azadiene and cyclopentadiene........................................................................................... 33 Figure 21. The Diels-Alder reaction of methyl vinyl ketone and 1,3-cyclohexadiene catalyzed by indium trichloride or methylrhenium trioxide. ............................................ 34 Figure 22. The induced fit mechanism for enzyme catalysis............................................ 37 Figure 23. The graphical definition of the Km and Vmax Parameters in the Michaelis Menten Equation............................................................................................................... 39 Figure 24. A cascade reaction involving o-quinones obtained by an enzyme-initiated hydroxylation-oxidation sequence combined with a Diels-Alder reaction....................... 43 Figure 25. Lipase catalyzed-domino reaction in the one-pot synthesis of optically active 7-oxabicyclo[2.2.1]heptenes (* represents chiral center). ................................................ 44 Figure 26. β-Glucosidase-triggered rearrangement of multifloroside in aqueous medium. ............................................................................................................................ 45 Figure 27. The synthesis of bicycle[3.1.0]hexane compound via enzyme-triggered Meinwald rearrangement. ................................................................................................. 46 Figure 28. Enzymatic dehydration-initiated Rearrangement of paclitaxel precursors ..... 47 Figure 29. Ester hydolysis-initiated dioxetane fragmentation. ......................................... 48 Figure 30. Enzymatic liberation of carboxylate anoin for the formation of γ-lactone ..... 49 Figure 31. Enzyme-initiated a tree-step SN2 cascade reaction of the diepoxide compound.......................................................................................................................... 50 Figure 32. Cyclisation of a diasteromeric mixture of (±)-epoxy ester initiated by enzymatic generated hydroxyl group................................................................................ 51
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Figure 33. Epoxide hydrolases initiated cyclisation of haloalkyl-oxiranes. ..................... 51 Figure 34. Enzyme-triggered transformation of meso-bis-epoxides................................. 52 Figure 35. Enzyme-catalyzed intramolecular 1,3-dipolar cycloaddition reaction............ 53 Figure 36. Two enzymetic reactions for the synthesis of cephalexin ............................... 54 Figure 37. Four enzyme system for domino synthesis of L-fructose. .............................. 55 Figure 38. Two enzyme system for the synthesis of enantiopure epoxide. ...................... 55 Figure 39. Active site of laccase CotA from Bacillus subtilis (adapted from Enguita et al.) .................................................................................................................... 58 Figure 40. Catalytic cycle of laccase showing the mechanism of four-electron reduction of a dioxygen molecule to water at the enzyme copper sites (adapted form Shleev et al. and Solomon et al.) ........................................................................................................... 61 Figure 41. Proposed decay mechanism of the native intermediate to the resting laccase. .............................................................................................................................. 62 Figure 42. (a) Scheme of laccase-catalyzed redox cycles for substrate oxidation; (b) The example of the oxidation of hydroquinone by laccase...................................................... 63 Figure 43. Chemical structure of laccase mediators. ........................................................ 65 Figure 44. Stucture of 3,3’,5,5’- tetramethoxy,1,1’-biphenyl-4,4’-diol produced by laccase catalyzed the oxidation of 2,6-dimethoxyphenol ................................................. 66 Figure 45. Dimer and tetramer products from the oxidation of isoeugenol alcohol by laccase. .............................................................................................................................. 67 Figure 46. Biotransformation of ferulic acid by laccase................................................... 68 Figure 47. The synthesis of bis-lactone lignans................................................................ 68 Figure 48. The oxidation of phenolic azo dyes by laccase ............................................... 69 Figure 49. The oxidative deprotection of p-methoxyphenyl (PMP)-protected amines by laccase. .............................................................................................................................. 73 Figure 50. The synthesis of actinocin by laccase mediated oxidation of 4-methyl-3-hydroxyanthranilic acid .................................................................................................... 74
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Figure 51. The synthesis of 2-amino-3H-phenoxazin-3-ones by the laccase catalyzed oxidative cycloaddition of o-aminophenols...................................................................... 74 Figure 52. The synthesis of the sulfonate analogue of cinnabarinic acid by laccase mediated the oxidative dimerization of 3-hydroxyorthanilic acid.................................... 75 Figure 53. The transformation of trans-resveratrol (3,5,4’-trihydroxystilbene) by laccase. .............................................................................................................................. 75 Figure 54. The oxidation of a seires of hydroxystilbenes by laccase ............................... 76 Figure 55. Laccase catalyzed the formation of catechin-hydroquinone adducts. ............. 78 Figure 56. Laccase catalyzed N-coupling of dihydrocaffeic acid and amines ................. 79 Figure 57. The synthesis of Tinuvin by a laccase-catalyzed reaction............................... 80 Figure 58. Mechanism of laccase mediated the formation of quinonoid intermediate for Michael addition reaction. ................................................................................................ 81 Figure 59. Laccase mediated amination reaction.............................................................. 82 Figure 60. The synthesis of 3-substituted-1,2,4-triazolo(4,3-β)(4,1,2)benzothiadiazine-8-ones by laccase mediated reaction of 5-substituted-4-amino-3-mercapto-1,2,4-triazoles and hydroquinone.............................................................................................................. 83 Figure 61. Laccase initiated domino reaction of cyclohexane-1,3-diones with catechols............................................................................................................................ 84 Figure 62. The synthesis of artificial urushi by laccase-catalyzed polymerization of urushiol analogues ............................................................................................................ 87 Figure 63. Structure of Rutin. ........................................................................................... 87 Figure 64. The structure of poly(8-hydroxyquinoline) ..................................................... 88 Figure 65. Chemical structure of cellulose ....................................................................... 90 Figure 66. Sugar monomers in hemicellulose................................................................... 92 Figure 67. Structure of hemicelluloses in softwood. ........................................................ 93 Figure 68. The structure of monolignols........................................................................... 94 Figure 69. Resonance structures of lignin precursors....................................................... 95
xviii
Figure 70. Structure of eight different lignin linkages...................................................... 95 Figure 71. Alkaline cleavage of α-aryl ether bond, sulfidolytic cleavage of β-aryl ether bonds in phenolic arylpropane units, and conversion into enol-ether units of quinone methide intermediates ....................................................................................................... 98 Figure 72. β-aryl ether bond cleavage in nonphenolic arylpropane unit .......................... 99 Figure 73. Competitive addition of external (SH-) and internal (phenolate ion) nucleophiles to quinone methide intermediates.............................................................. 100 Figure 74. Scheme illustrates peeling and stopping reactions of polysaccharides during kraft pulping.................................................................................................................... 102 Figure 75. A softwood tracheid (fiber) cell wall structure (Adapted from Coté). .......... 104 Figure 76. Laccase catalyzed grafting of lignin with 4-hydroxy-3-methoxybenzylurea, followed by chemical crosslinking to urea/formaldehyde (UF) resin in the subsequent glueing process................................................................................................................ 106 Figure 77. Proposed mechanism of chemoenymatically induced graft copolymerization between lignin and acrylamide. ...................................................................................... 108 Figure 78. Phenolic acids for the modification of high kappa pulp................................ 109 Figure 79. The proposed structure of the modified TMP with tyramine by laccase….. 111 Figure 80. Proposed mechanism for grafting of tyramine to lignin by laccase. ............. 112 Figure 81. Laccase catalyzed Coupling reaction of aminized cellulose with catechol... 113 Figure 82. Lipase-catalyzed reactions of triacylglycerols .............................................. 115 Figure 83. Examples of lipase-catalyzed reactions......................................................... 118 Figure 84. Asymmetric Michael addition reaction of 2-(trifluoromethyl)propenoic acid catalyzed by lipase from Candida cylindracea (* represents chiral center). .................. 120 Figure 85. Proposed mechanism of lipase catalyzed Michael addition of pyrrolidine and acrylonitrile. .................................................................................................................... 121 Figure 86. Michael addition of imidazole and methyl acrylate catalyzed by a variety of hydrolases ....................................................................................................................... 122 Figure 87. Michael addition of acetylacetone to acrolein catalyzed by a C. Antarctica lipase B Mutant ............................................................................................................... 123
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Figure 88. Graph illustrates the absorbance increase of laccase-oxidized ABTS at 420 nm. .................................................................................................................................. 125 Figure 89. Picture illustrates the changing in color of ABTS (in water) after adding laccase. The color changes from bright green to dark green. ......................................... 125 Figure 90. Photograph of the equipment set for soxhlet extraction................................ 126 Figure 91. Picture of Combiflash Companion instrument (Teledyne Isco company) with 40 g RediSep normal-phase silica flash columns ........................................................... 127 Figure 92. The reaction setting of the synthesis of 1,4-naphthoquinones and related structures via laccase-catalyzed Diels-Alder reaction. ................................................... 130 Figure 93. The titration data plotted as conductivity vs. volume of NaOH for the calculation of carboxyl group (RCOOH) content using conductivity method. .............. 134 Figure 94. Picture of instrument used for pulp disintegration. ...................................... 135 Figure 95. The PFI mill for the laboratory refining of pulp............................................ 136 Figure 96. Handsheet making apparatus (left) and handsheet made from liner board softwood kraft pulp (right).............................................................................................. 136 Figure 97. Tensile testers a) an Lorentzen and Wettre Alwetron tensile tester; b) an Instron tensile tester. ....................................................................................................... 137 Figure 98. An Elmendorf tearing tester. ......................................................................... 138 Figure 99. The Quinone Diels-Alder (QDA) reaction. ................................................... 142 Figure 100. The proposed reaction pathway of laccase-catalyzed Diels-Alder reaction of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a)............................... 143 Figure 101. 1H-MNR spectrum of crude mixture from the laccased-catalyzed reaction of of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a). Peaks of compound 3a are illustrated in blue boxes. Peaks of compound 4a are illustrated in red boxes. Peak of pentafluorobenzaldehyde is illustrated in green box. ................................................. 145 Figure 102. The preliminary reaction system for laccase-catalyzed aqueous Diels-Alder reaction of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a). ........... 153 Figure 103. The effect of laccase dose on the formation of compound 3a and 4a. The percent yield of 3a and 4a was measured by 1H-NMR spectroscopy. ............................ 155
xx
Figure 104. The effect of temperature on the formation of compound 3a and 4a. The percent yield of 3a and 4a was measured by 1H-NMR spectroscopy. (No products were obtained at 100 °C.) ........................................................................................................ 158 Figure 105. The example of enzyme-initiated reaction cascade reported by Waldmann and co-workers................................................................................................................ 164 Figure 106. Laccase-initiated cascade synthesis of substitute o-naphthoquinones via aqueous Diels-Alder reaction.......................................................................................... 165 Figure 107. The proposed mechanism for the elimination of methoxy or acetoxy from the reaction of 4-methyl-1,2-benzoquinone and 1-methoxy-1,3-butadiene or 1-acetoxy-1,3-butadiene in the presence of laccase in aqueous medium............................................... 178 Figure 108. Proposed mechanism of laccase/Sc(OTf)3 catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c). ............................................... 191 Figure 109. 1H-NMR of crude mixture from the laccase-catalyzed the reaction of 5a and 8a with and without lipase. These spectra demonstrate the formation of 9a and the decrease of starting material 5a during the reaction. ...................................................... 201 Figure 110. Proposed reaction pathway of laccase/lipase catalytic system for the synthesis of compound 9a............................................................................................................... 205 Figure 111. The formation of compound 9a from the reaction of 5a and 8a in the presence of laccase. The percent yield of 9a was measured by 1H-NMR spectroscopy.......................................................................................................................................... 209 Figure 112. The proposed mechanism of the elimination of Cl atom from the laccase/lipase catalyzed reaction of catechol and 8b in aqueous medium...................... 212 Figure 113. Proposed reaction pathway of laccase/lipase catalytic system for the reaction between catechol (5a) and aniline (10). .......................................................................... 215 Figure 114. Propose mechanism for the grafting treatment of high-lignin content softwood kraft pulp with laccase and amino acids. ........................................................ 221 Figure 115. (top) Bulk acid group content of control pulp, laccase treated pulp (Lac), glycine treated pulp (Gly), and laccase-glycine treated pulp (Lac/Gly) at different conditions (The control pulp, laccase treated pulp and Gly-treated pulp were treated in the same condition as laccase-Gly treated pulp but no laccase and Gly, no Gly, and no laccase, respectively); (bottom) bulk acid group content of pulps treated with laccase and different amount of glycine at pH 7.0 and room temperature......................................... 225
xxi
Figure 116. Bulk acid group content of (a) linerboard pulps treated with a variety of amino acids in the presence of laccase (80 U/g pulp); (b) linerboard pulps treated with a variety of amino acids (3.2 mmol/ 1g pulp) in the presence and absence of laccase. .... 229 Figure 117. Bulk acid group content of linerboard pulps that were treated with histidine (3.2 mmol/ 1g pulp) and different amount of laccase..................................................... 230 Figure 118. Nitrogen content of control pulp, lacccase treated pulp (Lac), and laccase-His treated pulp (Lac/His). .................................................................................................... 231 Figure 119. Physical paper properties of handsheets made from control pulp, laccase treated pulp (Lac), and laccase-histidine treated pulp (Lac/His); (a) tensile strength; (b) tear strength; (c) wet tensile strength. ............................................................................. 234 Figure 120. Scanning electron microscope (SEM) images of handsheets made from (a) control pulp; (b) laccase treated pulp; (c) laccase-histidine treated pulp........................ 235 Figure 121. Predictions from Page equation for tensile strength of paper vs. relative bonded area together with the qualitative effect of increasing fiber properties.............. 281 Figure 122. (a) The in-plane tear test; (b) the out-of-plane or Elmendorf tear test. ....... 285
xxii
LIST OF EQUATIONS
Page Equation 1. The Michaelis-Menten Equation (V=reaction velocity; Vmax = maximum reaction velocity; [S] = substrate concentration; Km = michaelis-menten constant; E = enzyme; S = substrate, P = product). 38 Equation 2. Lineweaver and Burk equation for determining Km and Vmax 40 Equation 3. The equation used to calculate for the carboxylic content of pulp fibers. 134 Equation 4. The Page equation. 280 Equation 5. Page’s equation for computing relative bonded area. 282 Equation 6. Parameters to plot for obtaining bond strength using the Page equation. 282
xxiii
NOMENCLATURE
4-HBA 4-Hydroxybenzoic acid
7-ADCA 7-Aminodesaacetoxycephalosporanic acid
ABTS 2,2’-Azinobis-(3-ethylbenzylthiozoline-6-sulphate)
Ala Alanine
Ar Aromatic
Arg Arginine
Asp Aspatic acid
ATP Adenosine triphosphate
DA Diles-Alder
DABCO 1,4-Diazabicyclo[2.2.2]octane
DCS Dodecanesulfonate
dDP 5,5-Di-n-dodecyl-2-hydroxy-1,3,2-dioxaphosphorinan-2-one
DMAP 4-Dimethylaminopyridine
DMSO Dimethylsulfoxide
DOPA 3,4-Dihydroxyphenylalanine
DP Degree of polymerization
DS Dodecylsulfate
E Enzyme
EC Enzyme commission
EPR Electron paramagnetic resonance
ES Enzyme-substrate complex
ESCA Electron spectroscopy for chemical analysis
ET Electron transfer
xxiv
EtOAc Ethyl acetate
FMO Frontier molecular orbital
FRPSG Fluorous reverse-phase siclica gel
FT-IR Fourier Transform Infrared
Gly Glycine
HAA Hydroxyanthanilic acid
HBT N-hydroxybenzotriazole
HCl Hydrochloric acid
His Histidine
HMBC Heteronuclear multiple bond coherence
HMQC Heteronuclear multiple quantum coherence
HOMO Highest occupied molecular orbital
Km Michaelis-Menten constant
Lac Laccase
LASCs Lewis acid/surfactant combined catalysts
Ln(OTf)3 Lanthanide triflate
LUMO Lowest unoccupied molecular orbital
MCD Magnetic circular dichorism
ML Middle lamella
M-M Michaelis Menten
MS Mass spectroscopy
NaCl Sodiun chloride
NaOH Sodium hydroxide
NMR Nucleae magnetic resonance
o.d. Oven dried
xxv
OH- Hydroxide anion
PAA 4-Hydroxyphenylacetic acid
PEG Polyethyleneglycol
Phe Phenylalanine
PMP p-Methoxyphenyl
QDA Quinone Diels-Alder
RT Room temperature
RTILs Room temperature ionic liquids
Sc(OTf)3 Scandium triflate
SDS Sodium dodecyl sulfate
SEM Scanning electron microscope
Ser Serine
SH- Hydrosulfide anion
T1 Copper atom type 1
T2 Copper atom type 2
T3 Copper atom type 3
TAPPI Technical association of the pulp and paper industry
TEMPO 2,2,6,6-tetramethyl-1-piperidinyloxyl
TLC Thin layer chromatography
TMP Thermomechanical pulp
UF Urea/formaldehyde
UV Ultraviolet
VA Violuric acid
Vmax Maximum reaction velocity
XAS Cu K-edge X-ray spectroscopy
xxvi
SUMMARY
Laccase (benzenediol:oxygen oxidoreductase, EC 1.10.3.2), a multi-copper-
containing oxidoreductase enzyme, is able to catalyze the oxidation of various low-
molecular weight compounds, specifically, phenols and anilines, while concomitantly
reducing molecular oxygen to water. Moreover, due to their high stability, selectivity for
phenolic substructures, and mild reaction conditions, laccases are attractive for fine
chemical synthesis. In this study, new green domino syntheses were developed by
conducting reactions in an aqueous medium, an environmentally-friendly solvent, and
using laccase as a biocatalyst.
The first study presents a work on the synthesis of naphthoquinones in the
aqueous medium. Herein, laccase was used to oxidize o- and p-benzenediols to generate
o- and p-benzoquinones in situ. These quinones then underwent Diels-Alder and
oxidation reactions to generate napthoquinone products. This reaction system can yield
naphthoquinones in up to 80% yield depending on the structure of the starting
hydroquinone and diene.
The next part of this thesis reports the cascade synthesis of benzofuran derivatives
from the reaction of catechols and 1,3-dicarbonyl compounds via oxidation-Michael
addition in the presence of laccase and Sc(OTf)3/SDS in an aqueous medium. Depending
on the substrates, one-pot yields of benzofurans averaged 50-79%. In the absence of
Sc(OTf)3, these yields decreased to 45-65%. Hence, the use of Lewis acid was critical for
efficient synthesis of the desired compounds. From an environmental concern, this
system still produced a hazardous waste from the transition metal catalyst. Therefore, the
xxvii
development of alternative methodologies to replace the lanthanide metal catalyst in this
synthesis is a high priority to enhance the overall green chemistry aspect. As a
consequence, lipase was used as a catalyst to replace Sc(OTf)3 for the synthesis of
benzofuran derivatives. The laccase/lipase co-catalytic system provides the benzofuran
products in a good yield. In addition, this catalytic system was also able to catalyze the
reaction of anilines and catechol.
Besides its application in organic synthesis, laccase also has an application in
fiber modification. Therefore, in the last part of this thesis, laccase was applied to the
modification of high-lignin softwood kraft pulp. This modification demonstrates the
potential of laccase-facilitated grafting of amino acids to high lignin content pulps to
improve their physical properties in paper products which resulted from the increase of
carboxylic acid group of the fibers. A unique two-stage laccase grafting protocol was
developed. Fibers were first treated with laccase, followed by grafting reactions with
amino acids. The bulk acid group content was measured, and a variety of amino acids,
including glycine, phenylalanine, serine, arginine, histidine, alanine, and aspartic acid,
were examined. The effects of laccase dosage and amino acids on fiber modification were
studied. In this study, histidine provided the best yield of acid groups on pulp fiber and
was used in the preparation of handsheets for physical strength testing. Laccase-histidine-
treated pulp showed an increase in the strength properties of the resulting paper.
1
CHAPTER 1
INTRODUCTION
1.1 Introduction
In recent years, the use of natural catalysts, enzymes, in the development of
organic synthesis reactions has received a steadily increasing amount of attention due to
their synthetic, economical, and, especially, environmental advantages [1,2]. The
enzymes are able to promote reactions under very mild conditions of temperature, pH,
and pressure. Moreover, to address the challenges of green chemistry, the possibility of
using water to replace the hazardous classical organic solvents in enzyme-catalyzed
reactions is another advantage. In addition to its environmental benefits, the use of water
as a solvent is both inexpensive and safe. The main purpose of this dissertation is to
create environmentally-friendly synthetic procedures by conducting the reactions in an
aqueous medium in the presence of a biocatalyst.
The main biocatalyst used in this dissertation is laccase. Laccase
(benzenediol:oxygen oxidoreductase, EC 1.10.3.2), a multi-copper-containing
oxidoreductase enzyme, is able to catalyze the oxidation of various low-molecular weight
compounds, including benzenediols, aminophenols, polyphenols, polyamines, and lignin-
related molecules, while concomitantly reducing molecular oxygen to water [3-10].
Because of its high stability, selectivity for phenolic substructures, and mild reaction
conditions, laccase is attractive for fine chemical synthesis [11-19]. Therefore, interest in
the potential use of laccase in organic synthesis has recently increased. Laccase also finds
a wide variety of industry applications, including food, pulp and paper, textile, cosmetics,
and nanobiotechnology industries [20,21]. Recently, laccase applications have shifted
toward fiber modification. Laccase has been reported to catalyze biografting of a variety
2
of substrates to technical lignins and lignin-rich cellulosic fibers [22-31]. Therefore, the
utilizing of laccase in green synthetic chemistry and in fiber modification was the main
focus of this research study.
In this dissertation, the synthesis of p-naphthoquinones and related structures via
Diels-Alder reaction of p-quinone generated by laccase and dienes in an aqueous media
was investigated. This study is described in Chapter 4. Chapter 5 further explores the
laccase-triggered Diels-Alder reaction of 1,2-hydroquinone and dienes for the synthesis
of o-naphthoquinones. Next, the cascade synthesis of benzofuran derivatives is
investigated in Chapter 6. This synthesis was conducted from the reaction of catechols
and 1,3-dicarbonyl compounds via oxidation-Michael addition in the presence of laccase
and Sc(OTf)3/SDS under air at room temperature in aqueous media. However, from an
environmental perspective, this system still produces a hazardous waste from the
transitional metal catalyst. Therefore, the development of alternative methodologies to
replace the lanthanide metal catalyst in this synthesis is a high priority in order to
enhance the overall green chemistry aspect of this one-pot synthetic reaction. As a
consequence, the enzyme named lipase was used as an alternative catalyst in conjunction
with laccase for the synthesis of benzofuran derivatives. In addition, this laccase/lipase
co-catalytic system was further investigated to catalyze the Michael addition of anilines
and catechols. The details of these studies are described in Chapter 7.
In addition, laccase also finds an application in fiber modification. In the last part
of this research study, Chapter 8, laccase was applied to the modification of high-lignin
softwood kraft pulp. This modification demonstrates the potential of laccase-facilitated
grafting of amino acids to high lignin content pulps to improve their physical properties
3
in paper products by increasing the carboxylic acid group of the fibers. Finally, some
overall conclusions and recommendations for future work complete the document.
1.2 Objectives
Recently, the increasing concern for the environment and for safe chemical
procedures requires the development of new green synthetic methods. Therefore, the
focus of this research is to develop new environmentally-friendly synthetic chemistry for
the synthesis of a wide variety of compounds. To address the challenges of green
chemistry, this study focuses on using a safer chemical, the enzyme laccase, in catalytic
amount, using an environmentally-benign solvent, water, and conducting the reaction at
ambient temperature. The major objectives of this research are summarized as follows:
Determine the potential use of laccase in organic synthesis
Develop new green chemistry synthesis by using a green reagent and a
green solvent, which are laccase and water, respectively.
Besides green synthetic applications, this study also investigated the application
of laccase in a new green procedure for modifying lignin-rich cellulosic fibers in an
aqueous medium. The major objectives of this fiber modification research are
summarized as follows:
Evaluate the feasibility of a system utilizing laccase to graft amino
acids with lignin-rich cellulosic fibers.
Develop a new green procedure for fiber modification.
4
Determine conditions where the laccase-facilitated grafting system is
the most effective for modifying fibers.
Evaluate the effect of laccase-facilitated grafting treatment on paper
strength properties.
5
CHAPTER 2
LITERATURE REVIEW
2.1 Green Chemistry
2.1.1 Definition of Green Chemistry
Green chemistry, also called sustainable chemistry, is a chemical philosophy
encouraging the design of products and processes that reduce or eliminate the use and
generation of hazardous substances. The U.S. Presidential Green Chemistry Challenge,
March 1995, defines green chemistry as,
“the use of chemistry for source reduction or pollution prevention, the highest tier
of the risk management hierarchy as described in the Pollution Act of 1990. More
specifically, green chemistry is the design of chemical products and processes that are
more environmentally benign”
2.1.2 Twelve Principles of Green Chemistry
Green chemistry is a highly effective approach to pollution prevention because it
applies innovative scientific solutions to real-world environmental situations. The 12
Principles of Green Chemistry, originally published by Paul Anastas and John Warner in
Green Chemistry: Theory and Practice [32]. These principles help to explain what the
definition means in practice. The principles cover such concepts as:
the design of processes to maximize the amount of raw material that ends up in
the product;
6
the use of safe, environment-benign substances, including solvents, whenever
possible;
the design of energy efficient processes;
the best form of waste disposal: do not create it in the first place.
The 12 principles are [32]:
1. Prevent waste: Design chemical syntheses to prevent waste, leaving no waste to
treat or clean up.
2. Design safer chemicals and products: Design chemical products to be fully
effective, yet have little or no toxicity.
3. Design less hazardous chemical syntheses: Design syntheses to use and generate
substances with little or no toxicity to humans and the environment.
4. Use renewable feedstock: Use raw materials and feedstock that are renewable
rather than depleting. Renewable feedstock are often made from agricultural
products or are the wastes of other processes; depleting feedstock are made from
fossil fuels (petroleum, natural gas, or coal) or are mined.
5. Use catalysts, not stoichiometric reagents: Minimize waste by using catalytic
reactions. Catalysts are used in small amounts and can carry out a single reaction
many times. They are preferable to stoichiometric reagents, which are used in
excess and work only once.
6. Avoid chemical derivatives: Avoid using blocking or protecting groups or any
temporary modifications if possible. Derivatives use additional reagents and
generate waste.
7
7. Maximize atom economy: Design syntheses so that the final product contains the
maximum proportion of the starting materials. There should be few, if any,
wasted atoms.
8. Use safer solvents and reaction conditions: Avoid using solvents, separation
agents, or other auxiliary chemicals. If these chemicals are necessary, use
innocuous chemicals. If a solvent is necessary, water is a good medium as well as
certain eco-friendly solvents that do not contribute to smog formation or destroy
the ozone.
9. Increase energy efficiency: Run chemical reactions at ambient temperature and
pressure whenever possible.
10. Design chemicals and products to degrade after use: Design chemical products to
break down to innocuous substances after use so that they do not accumulate in
the environment.
11. Analyze in real time to prevent pollution: Include in-process real-time monitoring
and control during syntheses to minimize or eliminate the formation of
byproducts.
12. Minimize the potential for accidents: Design chemicals and their forms (solid,
liquid, or gas) to minimize the potential for chemical accidents including
explosions, fires, and releases to the environment.
8
2.2 Water as Solvent in Organic Synthesis
In order to move toward sustainable technologies, developing more benign
synthetic procedures in chemical synthesis is important. This development can be
achieved by several approaches, including reducing the amount of waste, the energy
usage, and the use of volatile, toxic and flammable solvents. Therefore, many alternative
solvents have been proposed to replace classical organic solvents. The most well-known
of these alternate reaction media are listed below [33]:
Use of water as solvents
Reactions under solventless/solvent-free conditions
Supercritical carbon dioxide (31.1 ºC, 73 atm)
Supercritical water (374 ºC, 218 atm)
Room-temperature ionic liquids
Herein, the use of water as a reaction media is the main focus of this thesis. The
use of water as a medium for organic reaction is one of the finest solutions to the problem
of solvent toxicity and disposal. Water is the cheapest, safest and most non-toxic solvent
in the world. In addition, many surprising discoveries, such as an increase of reaction
rates and reaction selectivity, have been made when using water as a reaction medium.
The use of an aqueous medium affords both advantages and disadvantages, some of
which are listed below [34]:
Advantages:
Inflammable and anhydrous solvents are not needed
Economical saving
Abundant, cheap, not toxic and environmental friendly
9
Protection-deprotection of functional groups such as OH, COOH may not
be necessary
Water-soluble compounds can be used directly without derivatization
pH control
Preferred solvent for enzyme catalyzed reactions
Possibility of using additives such as mineral salts, surfactants,
cyclodextrins
Possibility of isolating products by decanting or filtration
Disadvantages:
Not inert
High boiling point
Problems isolating highly water-soluble products
Carbocarbon acid (pKa > 17) and water-sensitive reagents cannot be used
In the early 1980s, Breslow and Rideout were the first to show that Diels-Alder
reactions were greatly accelerated in water [35]. This discovery triggered a more
widespread interest toward the development of organic reaction in water. In the past 20-
30 years, the potential benefits of using aqueous media have been recognized, and
reactions including pericyclic, Michael additions, condensation, oxidation, reduction and
organometallic reactions have been reported [36-41]. Among the organic reactions
investigated in aqueous medium, the pericyclic reactions, especially Diels-Alder reaction,
has been the most widely studied [34,38,42,43]. The following section highlights some
Diels-Alder reactions that can be performed successfully in water.
10
2.2.1 Diels-Alder Reactions
The Diels-Alder reaction is a [4 + 2] cycloaddition in which a diene (4-π
component) reacts with a dienophile (2- π component) to provide a six-membered ring.
Bond-forming and bond-breaking processes are concerted in the six-membered transition
state (Figure 1). Most dienophiles are of the form –C=C−Ζ or Z−C=C−Z’, where Z and
Z’ are electron-withdrawing groups, such as CHO, COR, COOH, COCl, COAr, CN,
NO2, Ar, CH2OH, CH2Cl, CH2NH2, CH2CN, CH2COOH, halogen, PO(OEt)2, or C=C
[44]. Particularly common nucleophile are maleic anhydride and quinones. The Diels-
Alder reactions with quinones will be discuss in detail in the next section. When one or
more heteroatoms are present in the diene and/or dienophile framework, the
cycloaddition is called a hetero-Diels-Alder reaction. The Diels-Alder reaction is of great
value in synthetic organic chemistry because it creates the very useful cyclohexene ring.
Figure 1. Diels-Alder reaction of 1,3-butadiene with ethylene.
The reactivity, regiochemistry, and stereochemistry of the Diels-Alder reaction
can be explained by frontier molecular orbital theory (FMO). As applied to cycloaddition
reactions the rule is that reactions are allowed only when all overlaps between the highest
occupied molecular orbital (HOMO) of one component and the lowest unoccupied
+
Diene Dienophile
11
molecular orbital (LUMO) of the other are in phase such that a positive lobe overlaps
only with another positive lobe and a negative lobe only with another negative lobe.
These orbitals are the closest in energy [44]. Figure 2 illustrates the molecular orbitals of
alkenes and conjugated dienes, and the two dominant orbital interactions of symmetry
allowed Diels-Alder cycloaddition.
The reactivity of a Diels-Alder reaction depends on the energy difference between
HOMO and LUMO of the two components [43]. The lower the energy difference, the
lower is the transition state energy of the reaction. The energy level of both HOMO and
LUMO depends on the substituents. Electron-withdrawing groups lower their energy,
while electron donating groups increase their energy. For normal electron-demand Diels-
Alder reaction, the reaction is controlled by HOMO of diene and LUMO of dieneophile
(Figure 2). Therefore, the reactions are accelerated by electron-donating substituents in
the diene and by electron-withdrawing substituents in the dienophile. In contrast, the
inverse electron-demand Diels-Alder reaction is controlled by LUMO of diene and
HOMO of dienophile (Figure 2). Therefore, the reactions are accelerated by electron-
withdrawing groups in the diene and by electron-donating groups in the dienophlie.
12
Figure 2. Schematic drawing of the molecular orbitals of alkenes and conjugated dienes and the orbital interaction for normal and inverse electron demand Diels-Alder reactions.
Energy
LUMO
HOMOHOMO
LUMO
Z
HOMO
LUMO
Normal electron-demand Diels-Alder reaction
Energy
LUMO
HOMOHOMO
LUMO
Z
LUMO
HOMO
Inverse electron-demand Diels-Alder reaction
13
In addition, the regioselectivity of the Diels-Alder reaction can also be explained
by FMO theory. The regiochemistry is controlled by the orbital coefficients of the atoms
forming the σ-bonds. The σ-bonds form in such the way that the orbitals that have larger
coeffients (larger lobes in Figure 3) overlap together. The regioselective is increased
when the difference between the orbital coefficents of the two end atoms of diene and
two atoms of dienophile increase [43].
Figure 3. Example of the regioselectivity of normal electron-demand Diels-Alder reaction controlled by the orbital coefficients of the atoms forming the σ-bonds.[43] The FMO theory can be used to explain the stereochemistry of the Diels-Alder
reaction. The Diels-Alder reactions are suprafacial reactions and have two suprafacial
approached named endo and exo. In endo approach, the bulkier sides of diene and
dienophile lie one above the other. In exo approach, the bulkier side of one component is
under the small side of the other. Therefore, the exo addition mode is expected to be
preferred because of less steric repulsive interactions than in the endo approach.
EW
HOMO
LUMOED
ED
EW
+
ED
EW
EW = electron-withdrawing substituentED = electron-donating substituent
14
However, it appears that the endo adduct is usually the major product. This endo
preference can be explained by the FMO theory that the endo approach is kinetically
favored because of the additional nonbonding interaction called “secondary orbital
interaction” which stabilizes the endo transition state by lowering the trasition state
energy (Figure 4)[43]. This secondary orbital interaction can not be formed in the exo
approach.
Figure 4. The endo and exo approach of the Diels-Alder reaction between piperylene and acrolein and the secondary orbital interaction in the endo transition state.[43]
The main part of this dissertation focuses on the chemistry of quinonoid
compounds. Therfore, the next section will be discussed about the Diels-Alder reaction of
quinonoid compounds. Then, the Diels-Alder reactions carried out in the water under
conventional conditions of temperature and pressure will be illustrated next.
H3C
O
Secondary orbitalinteraction
endo
H3C
O
exo
No secondary orbitalinteraction
LUMO
HOMO
15
2.2.1.1 Quinone Diels-Alder Reaction
The quinone Diels-Alder (QDA) reaction (Figure 5) is a useful synthetic pathway
and many studies showed that the QDA adducts can be used as suitable starting points for
the synthesis of a wide variety of natural compounds, many of which are highly
functionalized.
Figure 5. The quinone Diels-Alder (QDA) reaction. An elegant example of the significance of QDA reactions in synthetic organic
chemistry was shown by R. B. Woodward in 1952. Woodward et al. created the route to
syntheis the steroids cortisone and cholesterol by using the QDA adduct of 2-methoxy-5-
methyl-p-benzoquinone and butadiene as a precursor for this synthesis. The bicyclic
adduct was formed via the intermediacy of endo transition state as illustrated in Figure 6.
O
O
O
O
16
Figure 6. A quinone-based Diels-Alder reaction as the key step in the total synthesis of the steroid hormones cortisone and cholesterol. Many studies have been reported the Diels-Alder reaction of quinonoid
compounds and several of these studies were reviewed by K. T. Finley [45]. Examples of
uncatalyzed and catalyzed quinone Diels-Alder reaction are summarized in Table 1.
O
O
MeO
Me
+
Benzene100 oC, 96h
86%
O
O
MeOMe
endo-transition state
O
O
Me
HMeO
aqNaOH, dioxanethen 1 Naq HCl
epimerization
O
O
Me
HMeO
Me
O
O
H
Me
H
OHOH
O
H
Me
HO
H
Me
H
H
H
Me
Me
Me
and
steps
Cortisone Cholesterol
17
Table 1. The examples of uncatalyzed and catalyzed quinone Diels-Alder reaction.
Reaction Reference
[46]
[47]
[48]
[49]
[50]
F
F
Ph3SiO
+
O
O
1. Benzene, 110 oC, 20 h45%
2. SiO2
OH
OHF
HO
NH
O
O
H
Δ
NH
H
O O
PhBr, Δ, 2.5h. 45%
NH
O
O
MeO
+
O
O
Toluene, ref lux
48 h, 57%
MeO
O
O
MeO
O
O
Me Me
+ Benzene, reflux24h, 82%
MeO
O
O
Me
H
H
Me
+
O
O
1. toluene, reflux, 20 h
2. DDQ, benzene, reflux, 5 h
O
O
Overall yield: 37%
18
Table 1. (Continued)
[51]
[52]
[53]
[54]
+
O
O
150 oC, 22 h
25%
O
O
+
O
O
Toluene, 100 oC
24 h, 79%
O
O
O
O
OCOMeO
O
Me Me
+100 oC, 0.5 h
Ethylene Glycol100%
O
O
MeMe
H
OCOMe
O
O
+K-10
0 oC, 5 h.70%
O
O
19
Table 1. (Continued)
[55]
[56]
[57]
The important application of quinone Diels-Alder reaction is to generate the QDA
adducts which can be used as the starting points in the total synthesis of various of natural
O
O
+10 mol% Sc(OTf)3
CH2Cl2, 0 oC
83%
O
O
O
O
OMe
O
+
Me
Et
10 mol% Catalyst
1:1 THF/toluene-78 oC
O
O
CO2Me
H
Me
Me
(98% ee, 87% yield)
NO
N N
O
SmH
PhPh
H
TfOOTf
OTf
Catalyst =
O
OOH
+
OCOMe
10 mol% catalyst, MS(-)
CH2Cl2, rt
O
OOH OCOMe
H
H
O
O
Ti
Cl
Cl
Catalyst =
20
compounds [58-60]. For example, a QDA reaction was used to construct the tricyclic
framework for the total synthesis of forskolin derivative [61]. The tricyclic carbon
skeleton of the analogue of forskolin was generated via a Diels-Alder cycloaddition
between a quinone and a vinyl cyclohexene as illustrated in Figure 7.
Figure 7. A Diels-Alder reaction of quinone and a vinyl cyclohexene as the key step in the total synthesis of forskolin derivative.[61] Recently, the Nicolaou group reported the use of Mikami’s catalyst ((S)-BINOL-
TiCl2) in the total synthesis of the unique terpenoid (-)-colombiasin A [62,63]. The first
step of this synthesis involved a selective asymmetric Diels-Alder reaction of
Danishefsky-type diene and quinone in the presence of the Mikami catalyst (30 mol%) as
shown in Figure 8. After many steps, (-)-colombiasin A was received in 32% overall
yield. White and Choi extended the versatility of this Mikami’s catalyst in their total
synthesis of (-)-ibogamine [64]. In this study, the Diels-Alder reaction of 1,4-
benoquinone and 1,3-diene catalyzed by Mikami’s catalyst was used as the key step in an
asymmetric synthesis leading to the alkaloid (-)-ibogamine (Figure 9). The preparation of
(-)-ibogamine was preceeded in 14 steps from 1,4-benzoquinone and the final product
was received in 10% overall yield.
OTBS
+
O
O
toluene
reflux60%
OTBSH
H
H
O
O Steps
OHH
OH
H
OEt
OAc
Forskolin derivative
21
Figure 8. A Diels-Alder reaction of Danishefsky-type diene and quinone in the presence of the Mikami’s catalyst for the total synthesis of (-)-colombiasin A.[62,63]
Figure 9. A Diels-Alder reaction of 1,3-diene and 1,4-benzoquinone in the presence of the Mikami’s catalyst as a key step for the total synthesis of ibogamine.[64]
Me
TBSO
+
O
O
OMe
Me
30 mol% (S)-BINOL-TiCl2
toluene, -60 − -10oC, 7 hO
O
Ti
O O
OMe Me
Me
OTBS
90%, 94%ee
Me
TBSO
O
O
OMe
Me
H
H
steps
Me
H
O
O
OMe
Me
MeMe
H
(-)-Colombiasin A32% overall yield
O
O
+
OTBS
TBS= tBuMe2Si
30mol%(S)-BINOL-TiCl2
CH2Cl2, rt, 0.5h
O
O
H
H
OTBS
steps
65%, 87%ee
NH
N
(-)-Ibogamine10%overall yield
22
Most recently, Jocobsen et al. [65] reported the application of the Cr-catalyzed
asymmetric quinone Diels-Alder Reaction for the total syntheses of (-)-colombiasin A
and (-)-elisapterosin B. The QDA adduct was used as a precursor for these syntheses. The
synthesis of (-)-colombiasin A was accomplished in 11.5% overall yield as summarized
in Figure 10.
Me
TESO
Me
+
O
O
OMe
Me
10 mol% catalyst
MS, toluene, 0 oC, 24 h
O
O
OMe
MeTESO
Me
MeH
H
O
O
OMe
MeTESO
Me
MeH
H
+
86% (combined)
steps
O
O
OMe
Me
HMe
H
Me
Me
Colombiasin A11.5% overall yield
BF3.Et2O, CH2Cl2
rt, 4hO
OH
O
Me
HMe
H
Me
Me
Elisapterosin B
94%
Me
ONCr
O(OH2)2Cl
Catalyst =
Figure 10. Cr-catalyzed asymmetric quinone Diels-Alder reaction as a key step for the total syntheses of (-)-colombiasin A and (-)-Elisapterosin B.[65]
2.2.1.2 Uncatalyzed Diels-Alder Reaction in Aqueous Medium
In 1931, Diels and Alder provided the first report of an uncatalyzed aqueous
Diels-Alder reaction of furan and maleic anhydride [66,67]. However, the first kinetic
study of acceleration of Diels-Alder reaction in water was studied by Rideout and
23
Breslow in 1980 [35]. In this study, they discovered that the reaction between
cyclopentadiene and methyl vinyl ketone in water was 740 times faster than in the apolar
hydrocarbon isooctane (Figure 11). By adding lithium chloride (salting-out agent) the
reaction rate increased 2.5 times further. The authors suggested that this unusual
acceleration in water was attributed to the polarity of the medium and hydrophobic
interaction (hydrophobic packing of diene and dienophile). The presence of lithium
chloride increased the reaction rate because the salt made the apolar reactants less soluble
in water and in so doing it enhanced the hydrophobic interaction.
Figure 11. Diels-Alder reaction between cyclopentadiene and methyl vinyl ketone in water and organic solvents.[35] Several experimental studies [68-71] and computer simulations [72] seem to
indicate that the rate enhancement of the aqueous Diels-Alder reactions are due to the
enforced hydrophobic interactions and hydrogen bonding interactions. The term
“enforced” is used to stress the fact that the association of the nonpolar reagents is driven
COMe
COMe
COMe+ 20-25 oC
endo exo
Isooctane 1Methanol 13Water 740Water + LiCl 1818
krel
Cyclopentadiene 3.9Ethanol 8.5Water 21.4
Endo/Exo
24
by the reaction and only enhanced by water. For instance, Engberts and his co-workers
[71] reported a kinetic study of a Diels-Alder reaction of 2,3-dimethyl-1,3-butadiene and
with N-methyl-, N-ethyl-, N-propyl-, and N-butylmaleimide in different solvents. These
reactions were accelerated in water relative to organic solvents as a result of enhanced
hydrogen bonding and enforced hydrophobic interactions during the activation process.
In addition, the acceleration increased as the hydrophobic character of the alkyl chain of
the dienophile increased (Figure 12).
Figure 12. Relative reaction rate (kwater/ kn-hexane) of Diels-Alder reaction between 2,3-dimethyl-1,3-butadiene and N-alkylmaleimides.[71]
Moreover, Sharpless and his colleagues [38] studied the cycloaddition of the
water insoluble trans,trans-2,4-hexadienyl acetate and N-propylmaleimide under various
conditions. The results of this study showed that the reaction in water suspension
provided substantial rate acceleration over homogeneous solution and the reaction in a
protic solvent such as methanol performed faster rate than in nonprotic solvent such as
acetonitrile and toluene (Figure 13). These results show that hydrogen bonding and
hydrophobic effects both are important for rate acceleration. Recently, Kumar and Tiwari
[73] explored three simple Diels-Alder reactions involving cyclopentadiene with methyl
N
O
O
RWater
25 oCN R
O
O
H
H
R krel
Me 1000Et 1447Pr 1683Bu 1881
+
25
acrylate, ethyl acrylate and butyl acrylate both in water and room temperature ionic
liquids (RTILs). They found that these Diels-Alder reaction in water are faster than in
RTILs. The reduction of reaction rate in RTILs can be attributed to the absence of
hydrophobic interactions and weaker hydrogen bonding in RTILs.
Figure 13. Diels-Alder reaction between trans,trans-2,4-hexadienyl acetate and N-propylmaleimide under various conditions.[38] Beside the rate enhancement, the enhancement of endo/exo selectivity of the
aqueous Diles-Alder reaction was also observed. Breslow et al. [74] also noted that the
endo addition of the reaction of cyclopentadiene with methyl vinyl ketone is more
favored when the reaction is carried out in water than when it is performed in organic
solvents (Figure 11). The endo preference in water were explained by the need to
minimize the transition state surface area in aqueous medium, thus favoring the more
compact endo transition state more than the extended exo transition state. Another
N(CH2)2CH3
O
O
AcO
+23 oC
N(CH2)CH3
H
H
O
O
AcO
Solvent Time (h) Yield (%)_________________________________
Toluene 144 79Acetonitrile >144 43Methanol 48 82Neat 10 82Water 8 81
26
example is the study of Grieco and his co-workers [75]. They examined the Diels-Alder
reaction between the α,β-unsaturated ketoaldehyde and ethyl 4-methyl-3,5-hexadienoate
(R = Et) in water and in hydrocarbon solvents (Figure 14). They found that the reaction
rate was doubled and both the reaction yield and the endo selectivity was enhanced when
conducting the reaction in aqueous medium. The best result was observed when
conducting the reaction of diene sodium carboxylate (R = Na). The reaction was
completed in 5 hours and the endo adduct is 75% of the diastereoisomeric reaction
mixture. In 1993, Paul et al. [76] applied this Diels-Alder reaction as a key step in the
synthesis of chaparrinone and other quassinoids (naturally occurring substances with
antileukemic activity). Recently, Utley et al. [77] reported the efficient formation of the
endo-Diels-Alder adducts of the reaction between ortho-quinodimethanes, generated
cathodically in aqueous electrolyte, and N-methylmaleimide.
Figure 14. Diels-Alder reaction between α,β-unsaturated ketoaldehyde and ethyl 4-methyl-3,5-hexadienoate.[75]
OMe
HO
CHO
CO2R
RT+
OMe
HO
CO2R
CHO
H
OMe
HO
CO2R
CHO
H
endo
exo
R Solvent time (h) endo/exo Yield (%)
Et PhH 288 0.85 52Et H2O 168 1.3 82Na H2O 5 3.0 100
27
Several studies have been reported the hetero Diels-Alder cycloadditions in
aqueous medium. For example, Kibayashi et al. explored the Diels-Alder reactions of the
nitroso moiety of the N-acylnitroso, a powerful dienophile, with a diene in water. The N-
acylnitroso compounds were generated in situ by periodate oxidation and then reacted
with dienes to form the Diels-Alder adducts. This N-acylnitroso compounds can be
trapped rapidly, especially in an intramolecular reaction such as the reaction of the in
situ-generated N-acylnitroso compound in Figure 15 that immediately cyclized to cis and
trans-1,2-oxainolactams [78]. Kibayashi et al. also used this acylnitroso approach in the
syntheses of (-)-swainsonine and (-)-pumiliotoxin [79].
Figure 15. Intramolecular hetero Diles-Alder reaction of N-acylnitroso compound.[78]
NHOH
O
OMOM
Pr4NIO4
0 oC, 1 minH2O
N
O
OMOM
O
H2O
NO
MOMO H
O
H
NO
MOMO H
O
H
+
trans-adduct
cis-adduct
trans/cis = 82:18
28
Lubineau and coworkers [80,81] have shown that glyoxylic acid, pyruvaldehyde,
and glyoxal were shown to react with cyclic or non-cyclic dienes via the aqueous hetero
Diels-Alder reaction to give the corresponding cycloadducts and/or α-hydroxy γ-lactones
in a good yield. Moreover, they also used this approach to prepare key starting
compounds for the enantioselective synthesis of 3-deoxy-D-manno-2-octulosonic acid
[82] and ketodeoxyheptulosonic acid derivatives [83]. Lubineau et al. have done the
extensive work in the studied of the aqueous Diels-Alder reactions to prepare optically
active oligosaccharides [84,85]. Some examples of Lubineau’s work are summarized in
Table 2. Another example for intramolecular hetero-Diels-Alder reaction in water was
reported by Grieco and Kaufman [86]. They examined the intramolecular Diels-Alder
reaction of iminium ions in polar media such as 5.0 M lithium perchlorate-diethyl ether
and water. In hot water, the tricyclic amine product can be obtained as the exclusive
diastereomer in 80% yield (Figure 16). They suggested that water appears to be the polar
solvent of choice for this reaction system because the use of lithium perchlorate-diethyl
ether as polar solvent led to some major problems. These problems occurred from the fact
that weak acid (lithium perchlorate) in highly polar media become strong acids and
protonation of the tethered dienes with concomitant diene isomerization is competitive
with cycloaddition.
29
Table 2. Example of the hetero Diels-Alder reactions studied by Lubineau et al.
Reactions Reference
[80]
[81]
[82]
[83]
[85]
+H COOH
O
H2O, pH 140 oC, 1.5 h
O
H
H
H
O +
O
H
H
H
O
73:2783%
+H COOH
O
100 oC, 1.5 h97%
H2O O
COOH
O
COOH
+
64:36
+H COONa
O
HO
HO
3. Ac2O/pyridine54%
1. H2O2. MeOH/H+ O
CO2CH3
AcO
AcO
O
HO
HO
COOH
OH
HO
OH
+H COONa
O
HO
1. H2O, 100 oC
2. H+/CH2N2
O
CO2CH3
OH
OCO2CH3
OCH3
AcO
OAc
OH
OHO
HOOH
H
OH
+H COONa
O 1. H2O, 140 oC, 48 h2. MeOH, Dowex-50 (H+)3. Ac2O-pyridine
68%
OAcO
AcO
OAc
H
OAc
O CO2Me
O
OH
HOHO
HOO
OH
30
Figure 16. Intramolecular imino-Diels-Alder reactions.[86]
2.2.1.3 Lewis-Acid-Catalyzed Diels-Alder Reaction in Aqueous Medium
In recent years, a number of water-tolerant Lewis acids have been used to catalyze
various Diels-Alder reactiond in aqueous medium [34]. In 1993, Kobayashi [55] reported
the use of scandium trifate, Sc(OTf)3 for the Diels-Alder reaction in aqueous medium.
This catalyst was stable in water and easily recovered to reused. Many other Lewis acids
have been reported to catalyze Diles-Alder reactions in water. Engberts [87,88] reported
the use of aqua-complexing agents including Co(NO3)2.6H2O, Ni(NO3)2.6H2O,
Cu(NO3)2.3H2O, and Zn(NO3)2.4H2O as Lewis acid catalysts for Diels-Alder reaction in
aqueous medium. The Diels-Alder reactions performing in aqueous medium in the
presence of these metal catalysts were faster than the aqueous reactions without the
catalysts, and Cu2+ ion showed to be the best catalyst in this study. However, the catalysts
worked efficiently only if they formed a chelate with the dienophile, and complexation
with α-amino acids (see Figure 17) which induces asymmetry in the Diels-Alder reaction
as in the copper-catalyed the reaction of 3-phenyl-1-(2-pyridyl)-2-propen-1-one with
cyclopentadiene (Figure 18) [89]. This cycloaddition occurs endo-stereoselectively in 3
days with high yield and with acceptable enantioselectivity (ee = 74%). Therefore, this is
NTFA NH
H
80%
H2O, 70 oC
31
the first enantioselective Lewis acid-cataltzed Diels-Alder reaction in water. Recently,
Engberts and Mubofu [90] reported a comparative study of specific acid catalysis
(hydrochloric acid) and Lewis acid (i.e. copper (II) nitrate) catalysis of Diels–Alder
reactions in aqueous medium. They found that the reaction rate is 40 times faster with
copper catalysis than with hydrochloric acid catalysis at equimolar amounts of copper(II)
nitrate and hydrochloric acid and under the same reaction conditions.
Figure 17. Complexation of Cu(L-abrine) catalyst and 3-phenyl-1-(2-pyridyl)-2-propen-1-one.[89]
Figure 18. The enantioselectivity of copper (L-arabine) catalyzed Diels-Alder reactions of 3-phenyl-1-(2-pyridyl)-2-propen-1-one with cyclopentadiene.[89]
N
O
O
N
O
NCat. Cu(L-abrine)0 oC
+
+
endo exo
Solvent ee (%)H2O 74EtOH 39CH3CN 17THF 24CHCl3 44
N
O
O
NH
H3C
Cu2+O
N
R
R =
32
Many studies have now used water-tolerant Lewis acid, lanthanide triflates
(Ln(OTf)3) [91] together with Bi(OTf)3 [92], Sc(OTf)3 [93] and In(OTf)3 [94,95] to
catalyze the Diels-Alder reactions in water. For example, Wang et al. [96] studied the use
of Ln(OTf)3 to catalyze the aqueous aza-Diels-Alder reaction of an aldehyde and amine
hydrochloride with diene. Figure 19 shows a representative reaction of this study. The
product (endo + exo) was isolated in only 4% yield when no Ln(OTf)3 was added.
However, the yield of the product was increased to 64% when the lanthanide catalyst was
added.
Figure 19. The aqueous aza-Diels-Alder reaction using lanthanide triflate.[96] Lanthanide triflates were also shown to catalyze imino Diels-Alder reactions of
imines with dienes or alkenes which were developed by Kobayashi and his co-workers
[97]. Here, they reported a three-component coupling reactions between aldehydes,
amines, and dienes or alkenes which were successfully carried out by using lanthanide
triflate as a catalyst to afford pyridine and quinoline derivatives in high yields (Figure
20). Recently, Taguchi et al. [95] developed indium(III) triflate catalyzed intramolecular
Diels-Alder reaction of ester-tethered 1,7,9-decatrienoates in aqueous media. This
reaction gave the cycloadducts in good yield with perfect endo-selectivity and In(OTf)3 is
recyclable without troublesome purification.
H
O
NBn
NBn
exo endo
Ln(OTf)3, H2O+ + +BnNH3Cl
33
Figure 20. Yb(OTf)3-catalyzed Diels-Alder reaction between N-benzylideneaniline as azadiene and cyclopentadiene.[97]
Lewis acid/surfactant combined catalysts (LASCs) such as M(DS)n, M(DCS)n,
[98,99] and Cu(dDP)2 [100] (M = lanthanides, Sc, Yb, Cu, Zn, Ag, Mn, Co; n = 1, 2, 3;
DS = dodecylsulfate, DCS = dodecanesulfonate, dDP = 5,5-di-n-dodecyl-2-hydroxy-
1,3,2-dioxaphosphorinan-2-one) have recently been prepared. However, reports on their
catalytic ability in Diels-Alder reactions are discrepant.
Indium trichloride [101,102] and methylrhenium trioxide [103] are also water-
tolerant Lewis acids, and have been reported to catalyze Diels-Alder cycloadditions in
water. Some examples of these catalyst in the cycloaddition of methyl vinyl ketone and
1,3-cyclohexadiene are illustrated in Figure 21.
HCHO +
NH2
Cl
N
Cl
CH2
NH
H
ClH
H2O-EtOH-PhH(1:9:4)
Yb(OTf)390%
34
Figure 21. The Diels-Alder reaction of methyl vinyl ketone and 1,3-cyclohexadiene catalyzed by indium trichloride or methylrhenium trioxide.[101,103] Recently, Nishikido et al. [104] reported fluorous reverse-phase silica gel
(FRPSG)-supported Lewis acids catalyzed Diels-Alder reactions in water, and the
FRPSG-supported Lewis acids could be recycled by simple filtration after the reaction.
Yu et al. [105] examined the use of water-soluble organotungsten Lewis acid, [OP(2-
py)3W(CO)(NO)2](BF4)2 to catalyze Diels-Alder reactions under conventional heating or
microwave heating conditions. The cycloaddition reactions were efficiently conducted in
either water or in an ionic liquid, 1-butyl-3-methylimidazolium hexafluorophosphate.
Most recently, Litz [106] reported Flextyl PTM, a novel Ti(IV) performance catalyst,
catalyzed the aqueous Diels-Alder reaction of 1,3-cyclohaxadiene with 1,4-
benzoquinone. The catalyst improved conversion by 22% versus the uncatalyzed
reaction.
O
Water, r.t.
O
+ +
O
endo exoendo/exo Yield (%)
InCl3 90:10 87MeReO3 99:1 91
Catalyst
35
2.3 Biocatalysis
2.3.1 Enzymes
Enzymes are natural catalysts that accelerate the rate of reactions. Like all
catalysts, enzymes work by lowering the activation energy (Ea or ΔG‡) for a reaction,
thus dramatically increasing the rate of the reaction. Enzymes are composed of one or
more polypeptides organized in a specific three-dimensional structure through
interactions between the functional groups on the amino acid constituents. These
interactions include ionic bonding, covalent bonding, hydrogen bonding, and van der
waal’s forces. Some of the outstanding features of the enzymes include high substrate
specificity, specificity in promoting only one biochemical reaction with their substrate
ensuring synthesis of a specific biomolecular product without the concomitant production
of by products, stereospecificity, and regeospecificity, which they express in catalysis.
2.3.1.1 Nomenclature and Classification
An enzyme’s name is often derived from its substrate or the chemical reaction it
catalyzes, with the word ending in “ase”. For identification purpose, the International
Union of Biochemistry and Molecular Biology have developed a nomenclature for the
enzymes. Every enzyme has a four-digit number in the general form EC A.B.C.D, where
EC stands for ‘Enzyme Commission’; the following properties are encoded:
A indicates to which of the six main divisions (classes) the enzyme belongs,
B stands for the subclass, indicating the substrate class or the type of transferred
molecule,
C indicates the nature of the co-substrate,
36
D is the individual enzyme number.
Enzymes have been classified into six categories according to the type of reaction
they catalyze. These six classes of enzymes are listed below:
Class 1 – Oxidoreductases: catalyze oxidation/reduction reactions,
Class 2 – Transferases: transfer a functional group such as methyl or phosphate
group,
Class 3 – Hydrolases: catalyze the hydrolysis of C-O, C-N, O-P and C-S bonds,
Class 4 – Lyases: catalyze the addition or removal of some chemical groups of
substrate by mechanism other than oxidation, reduction, or hydrolysis,
Class 5 – Isomerases: catalyze isomerization changes within a single molecule,
Class 6 – Ligases: catalyze the joining together of two compounds coupled with
the hydrolysis of a diphosphate bond in ATP or a similar triphosphate.
2.3.1.2 Enzyme Mechanism
Enzymes are three-dimensional proteins that possess an “active site”. At the
active site, specific amino acids interact with the substrate, and the tranfornation of
substrate take places. In order to understand enzyme catalysis, some models have been
proposed.
2.3.1.2.1 ‘Lock-and-Key’ Mechanism
In 1894, Emil Fischer [107] developed the first proposal for a general mechanism
of enzymatic action. He hypothesized that an enzyme and its substrate form a complex
very much like a “lock and key”; therefore, each enzyme is very substrate specific and its
structure is completely rigid. However, this model can not explain why many enzymes do
37
act on large substrates, while they are inactive on smaller counterparts. Moreover, this
hypothesis can not explain why many enzymes can convert a variety of nonnatural
compounds besides their natural substrates [108]. Thus, another model had to be
developed.
2.3.1.2.2 Induced-Fit Mechanism
Daniel Koshland [109] suggested a modification to the lock and key model that
the enzymes are not entirely rigid but rather represent delicate and soft structures. During
the formation of the enzyme-substrate complex, the enzyme can change its conformation
under the influence of the substrate structure so as to wrap itself around its guest (Figure
22). This phenomenon was denoted as the ‘Induced Fit’. The induced fit theory states a)
precise orientation of catalytic groups is required for enzyme action b) the substrate
causes changes in the amino acids at the active site c) the changes in the catalytic
structure caused by a substrate will bring the catalytic groups into proper alignment
whereas a non-substrate will not achieve this.
Figure 22. The induced fit mechanism for enzyme catalysis.
Enzyme Substrate
Active Site
Enzyme Substrate
The active site changes its shapeto better fit the substrate.
38
2.3.1.3 Enzyme Kinetics
The rate at which an enzyme converts substrate to products is referred to as its
“activity”. When a smaller amount of enzyme can convert a greater amount of substrate
it is said to be more “active”. The reaction kinetics have been characterized for many
enzymes. Enzymatic activity is the productivity of the enzyme defined under strict
standard conditions. Michaelis and Menten [110] used a simple unimolecular reaction to
extract relationships used for predicting the kinetic properties of enzymes (Equation 1).
The symbols that describe the reaction are E=Enzyme and S=Substrate. The reaction
described by Michaelis and Menten proceeds in three phases. The initial or stationary
phase is an important phase as it is at this point where substrate and enzyme come
together for the intimate contact at the enzyme active site for the reaction. The second
phase of the enzyme reaction is the steady state where the enzyme is assumed to be
completely saturated with substrate and the rate of the reaction is dependant on the
amount of enzyme (E) or enzyme-substrate complex (ES). According to Michaelis
Menten (M-M) kinetics, the rate-limiting step is the conversion from ES to the product
(P). The Michaelis Menten relationship is stated in Equation 1.
V= Vmax [S]/[S]+Km
E + S ES E + P
Equation 1. The Michaelis-Menten Equation (V=reaction velocity; Vmax = maximum reaction velocity; [S] = substrate concentration; Km = michaelis-menten constant; E = enzyme; S = substrate, P = product).
k1
k2
k3
39
Km is the M-M constant and k3 is the turnover constant. These factors are
important for gauging the efficiency of an enzyme-substrate system. Km is the
concentration of substrate required for an enzyme to reach one-half of its maximum
velocity or Vmax. Essentially, Km is an indicator of the sensitivity or affinity of a
particular enzyme for a certain substrate (Figure 23).
Figure 23. The graphical definition of the Km and Vmax Parameters in the Michaelis Menten Equation
The turnover number is the rate at which the enzyme-substrate complex is
converted to the product, which indicates the ability of the enzyme to convert substrate
into product. Since k3 is the rate of formation of the product and Km is the affinity of the
enzyme for the reactants, the value k3/Km is usually a measure of the total enzyme
productivity [111], therefore, achieving a maximum velocity at a low substrate
concentration is ideal. Eventually the substrate concentration becomes limiting, and the
reaction reaches its asymptotic limit [111] (Figure 23). Kinetic units can be elucidated by
a relationship derived by Lineweaver and Burk (Equation 2).
40
1/V=1/Vmax+Km/Vmax x1/[S]
Equation 2. Lineweaver and Burk equation for determining Km and Vmax
Plotting the reciprocal of reaction rate vs. the reciprocal substrate concentration
allows one to obtain 1/Vmax at the y-intercept and -1/Km at the x-intercept.
2.3.1.4 Advantages and Disadvantages of Biocatalyst
2.3.1.4.1 Advantages of Biocatalysts [108]
Enzymes are very efficient catalysts: Compare to the nonenzymatic reactions, the
rates of enzyme-mediated processes are accelerated by a factor of 108-1010.
Enzymes are environmentally benign reagents.
Enzymes act under mild conditions: Enzymes act in a range of about pH 5-8, and
in a temperature range of 20-40 ºC. This minimizes problems of undesired side
reactions. However, there are some thermostable enzymes that can be performed
at high temperature.
Enzymes are compatible with each other: Several biocatalytic reactions can be
carried out in a reaction cascade in one reactor because enzymes normally
function under the same or similar conditions.
Enzymes are not bound to their natural role: Enzymes can catalyze a variety of
nonnatural substrates and often they are not required to work in water.
Enzymes can catalyze a broad spectrum of reactions.
Enzymes display selectivity: Three major types of selectivity are chemoselectivity,
regioselectivity and diastereoselectivity, and enantioselectivity.
Valuable resource for green chemistry
41
2.3.1.4.2 Disadvantages of Biocatalysts [108]
Enzymes are provided by nature in only one enantiomeric form.
Enzymes require narrow operation parameters: If a reaction proceeds too slow
under given parameter of temperature and pH, there is only a narrow operational
window for alteration. High temperature and extreme pH lead to deactivation of
the enzymes.
Enzymes display their highest catalytic activity in water.
Some Enzymes are bound to their natural cofactors such as NAD(P)H, and
chemical energy (ATP) : These cofactors are relatively unstable molecules and
are prohibitively expensive to use in stoichiometric amounts.
Enzymes are prone to inhibition phenomena: Many enzymatic reactions are
prone to substrate- or product-inhibition, which causes the enzyme to cease to
work at higher substrate and/or product concentrations, a factor which limits the
efficiency of the process.
Enzymes may cause auto-immune responses including allergies
2.3.2 Enzymes in Domino Reactions
Domino or cascade reactions involve two or more bond-forming transformations,
which take place under the same reaction conditions, without adding additional reagents
and catalysts, and in which the subsequent reactions result as a consequence of the
functionality formed by bond formation or fragmentation in the previous step all
occurring in one-pot [112]. The domino reaction is often proceeded via highly reactive
intermediates.
42
In recent years, the availability of enzymes has increased. Therefore, the use of
enzymes in the development of domino reaction has also increased in address the
challenges of Green Chemistry. Many studies involve enzyme-initiated domino reactions
have been reported [1,113-115]. Emzyme-initiated domino reactions follow a common
reaction sequence. Firstly, the enzyme modifies a group (‘trigger’ group) in the starting
material, generating a reactive intermediate that can undergo a subsequent domino
reaction consisting of a (i) fragmentation, (ii) rearrangement, (iii) cyclization such as
Diels-Alder reaction, or (iv) an intramolecular substitution affecting cyclization.
2.3.2.1 Enzyme-Triggered Diels-Alder Reaction
The first successful combination of an enzymatic with a nonenzymatic
transformation within a domino process was reported by Waldmann et al. in 1996
[116,117]. They reported the synthesis of highly functionalized bicycle[2.2.2]octenes by
a tyrosinase-initiated hydroxylation-oxidation of phenols followed by a Diels-Alder (DA)
reaction with electron rich dienophiles (see Figure 24). These studies, conducted in
chloroform in the presence of oxygen, provided a unique three-step one-pot reaction of
bicyclic DA products in high yields with the key intermediate being reactive ortho-
quinones.
43
Figure 24. A cascade reaction involving o-quinones obtained by an enzyme-initiated hydroxylation-oxidation sequence combined with a Diels-Alder reaction.[116,117] Kita and his co-worker [118,119] reported the first one-pot synthesis of optically
active 7-oxabicyclo[2.2.1]heptenes catalyzed by lipase, the hydrolase enzyme that act on
carboxylic ester bonds. As illustrated in Figure 25, the first step of this reaction was the
kinetic resolution of racemic furfuryl alcohol derivatives via acyl transfer catalyzed by
lipase. The next step was the intramolecular Diels-Alder reaction of the intermediate to
provide 7-oxabicyclo[2.2.1]heptene derivatives. Most recently these authors reported the
use of a lipase and a ruthenium catalyst to prepare polysubstituted decalines with high
optical and chemical yields from racemic alcohols [120].
OH
R1
OH
OH
R1
O
O
R1
Tyrosinase
CHCl3, O2
TyrosinaseCHCl3, O2
R2
O
O
R1R2
O
O
R1R2
+R2 = OEt, Ph
R1 = H, Me, iPr, tBu F, Cl, Br, I, OMe
Yield: 51-77%
44
Figure 25. Lipase catalyzed-domino reaction in the one-pot synthesis of optically active 7-oxabicyclo[2.2.1]heptenes (* represents chiral center).[118,119] 2.3.2.2 Enzyme-Triggered Rearrangement
Skeleton rearrangements are a special class of reactions in organic synthesis
because they often lead to products of exceptional structure. β-Glucosidase has been
reported to initiate rearrangement of multifloroside by the Shen group [121].
Multifloroside was subjected to β-glucosidase in acetate buffer. The domino process
started by enzymatic cleavage of a glycoside, and then a rearrangement subsequently
took place to generate jasmolactone analogues as the final products in a rather low yield
(10-20%) (Figure 26).
O
R2
OH
R3
R1
OEtOR4
OO
R2
O
R3
R1
R4
O
*
OO
O
R3R2
R1
R4
Lipase
Kinetic Resolution
Diels-Alder**
**
*
45
Figure 26. β-Glucosidase-triggered rearrangement of multifloroside in aqueous medium.[121]
An unusual enzyme-triggered asymmetric rearrangement was observed by Ohno
and his co-workers when they attempted to hydrolyze the asymmetric tricyclic diester in
an asymmetric fashion using porcine liver esterase [122]. First, a hemiester was form by
hydrolysis and then immediately underwent a Meinwald rearrangement to furnish the
final bicycle[3.1.0]hexane framework (Figure 27).
O
CO2R1
R2O
O
HOOGlu
H
β-GlucosidaseAcetate Buf fer O
CO2R1
H
OH
O
O
OR2
R1 = CH3, DHPR2 = DHP, H, CH3
10-20%
DHP =
OH
OH
Multifloroside
46
Figure 27. The synthesis of bicycle[3.1.0]hexane compound via enzyme-triggered Meinwald rearrangement.[122] During the development of a new method for the synthesis of paclitaxel, an
unexpected enzymatic dehydration-initiated rearrangement was discovered by Kim et al.
[123]. The 7-triehtylsilyl derivative of 10-deacetylbaccatine III served as a precursor for
this cascade reaction (Figure 28). In the presence of trichloroacetic anhydride as the acyl
donor, this precursor was acylated by Rhizopus delemar lipase at the 13-hydroxy group,
and underwent the dehydration-rearrangement to form the tricyclic diterpene
intermediate. After a prolonged reaction time, the intermediate underwent a second
dehydration to form the final product.
O CO2Me
CO2Me
Pig Liver Esterase
buffer pH 8, 30 oC
O CO2
CO2Me
HOCO2
CO2Me
OHC
CO2Me
CO2H
100%, ee = 48%
47
Figure 28. Enzymatic dehydration-initiated Rearrangement of paclitaxel precursors.[123] 2.3.2.3 Enzyme-Triggered Fragmentation
The Schaap group [124] presented the use of aryl esterase to catalyze the cleavage
of a naphthyl acetate-substituted dioxetane in aqueous buffer at ambient temperature. The
1,2-dioxetane moiety of the naphtyl acetate was cleaved via hydrolysis by porcine liver
esterase, thus generating the free intermediate naphtholate anion which subsequently
underwent fragmentation reaction to form the naphthol methyl ester and admantone with
the concurrent chemiluminescence (Figure 29).
OHO
HO
OCH2PhH
HO O
OSiEt3
OAc
13
OOHO
Et3SiO
OAc
OCH2Ph
O
Rhizopus delemar lipase(Cl3CCO)2O
- H2O
OO
Et3SiO
OAc
OCH2Ph
O
- H2O
48
Figure 29. Ester hydolysis-initiated dioxetane fragmentation.[124]
During the synthesis of N-Ras lipopeptides, Waldmann et al. [125] developed a
new protecting group for amino, hydroxyl, and carboxy moieties containing a p-
acetoxybenzyloxycarbonyl group. In this study, lipase was first used to cleave the acetate
group of the p-acetoxybenzyloxycarbonyl moiety to liberate the phenolate anion. Then,
this intermediate anion underwent a fragmentation to generate a quinone methide with
liberation of the desired products. This strategy was also applicable to solid-phase
synthesis. The aromatic moiety that was to build the scaffold was linked on to a
macroscopic polymeric carrier via a spacer-arm which acted as an enzymatically labile
anchoring group [126]. This method is useful for combinatorial chemistry and parallel
synthesis for the production of compound libraries attached to polymeric supports.
2.3.2.4 Enzyme-Triggered Intramolecular Substitution Affecting Cyclization
Enzyme-triggered intramolecular substitution affecting Cyclization reactions
normally start with the enzymatic hydrolysis of an ester or epoxide to form the
O O
OCOCH3
OCH3O O
O
OCH3
O
O
H3CO
Porcine Liver Esterase
+ Adamantanone + Light
49
hydroxylate or hydroxyl group which acts as a nucleophile to attack an electrophile via
intramolecular SN2 reaction in the second step. As in the work of Tamm et al., they
conducted the asymmetric hydrolysis of meso-epoxy diester using porcine liver esterase
in aqueous medium (Figure 30) [127]. In this cascade reaction, carboxylate anion was
liberated by enzymatic hydrolysis of the more accessible (equatorial) carboxy ester. This
carboxylate anion acted as nucleophile and attacked the epoxide moiety to generate the
corresponding hydroxyl γ-lactone. Due to a conformation change of the intermediate
during lactone formation, the remaining axial ester moiety was converted into the more
accessible equatorial ester which could be additionally hydrolyzed by the esterase. This
led to the formation of the final chiral product in 96% ee.
Figure 30. Enzymatic liberation of carboxylate anoin for the formation of γ-lactone.[127]
O
O
OCH3
O
OCH3
Pig Liver Esterasebuf fer pH 7
O
O
O O
OCH3
O
O
HO
OCH3
O
Pig Liver Esterasebuf fer pH 7
O
O
HO
OH
O
72% yieldee = 96%
50
Another example of enzymatic hydrolysis of ester to liberate the carboxylate
anion was reported by Williams et al [128]. In this study, the diepoxide underwent bis-
cyclization by the pig liver esterase, with stereospecific opening of each epoxide ring in a
5-exo-tet manner to form the final product. The reaction mechanism is summarized in
Figure 31.
Figure 31. Enzyme-initiated a tree-step SN2 cascade reaction of the diepoxide compound.[128] An alcoholic group generated from the enzymatic hydrolysis of ester or epoxide
can also act as nucleophile in a cascade reaction. For example, the ester moiety of a
diasteromeric mixture of (±)-epoxy ester was hydrolyzed by a crude immobilized enzyme
preparation (NOVO SP 409), or whole lyophilized cells of Rhodococcus reythropolis
NCIMB 11540 to generate the corresponding intermediate alcohol (Figure 32). The
alcohol immediately opened the epoxide in an SN2 fashion to furnish the corresponding
diastereomeric tetrahydrofuran derivatives [129].
O
OCH3
O O
H
O
O
O
O O
H
O
O O
H
O
OO
O
H
O
OO
OO
OO O
OH
PigLiver Esterase
phosphatebuffer pH7.5-8
70%
51
Figure 32. Cyclisation of a diasteromeric mixture of (±)-epoxy ester initiated by enzymatic generated hydroxyl group.[129] In the following example, the diol nucleophile was generated by emzymatic
hydrolysis of an epoxide to initiate a cascade reaction. For instance, the biohydrolysis of
(±)-2,3-disubstituted cis-chloroalkyl-epoxides (Figure 33) [130]. First, bacterial epoxide
hydrolases (Mycobacterium paraffinicum NCIMB 10420) hydrolyzed the racemic
epoxide to form the corresponding diol which underwent spontaneous ring closure to
yield the final cyclic product. This synthetic strategy has been used in asymmetric
synthesis of many bioactive compounds [131-133].
Figure 33. Epoxide hydrolases-initiated cyclisation of haloalkyl-oxiranes.[130]
O
O t-Bu
O
O
OH O
OH
O
OH
Rhodococcus sp.NCIMB 11540
Tris-buffer pH 7
70%
+
ee > 98% ee > 98%
1 : 1.2
O
n-Bu
Cl
n-Bu
Cl
OH
OHOH
n-BuOBacterial Epoxide Hydrolases
ee = 92%
O
n-Bu n-Bu
OH
OH
Bacterial Epoxide Hydrolases
ee = 86%Cl
(d,l)
(d,l)
ClO
OH
n-Bu
52
The enzyme triggered cyclisation of bis-epoxides using bacterial epoxide
hydrolase was investigated by Faber and his co-workers [134]. In this study, the
tetrahydrofuran products were generated through two secondary pathways as illustrated
in Figure 34. The products contain four stereogenic centers which constitute the central
core of bioactive Annonaceous acetogenins.
Figure 34. Enzyme-triggered transformation of meso-bis-epoxides.[134] 2.3.2.5 Enzyme-Triggered Other Type of Reactions
In 2005, the Kita group [135] developed a lipase-catalyzed domino kinetic
resolution of α-hydroxynitrone intramolecular 1,3-dipolar cycloaddition reactions which
successfully applied in the asymmetric total synthesis of (-)-rosmarinecine (Figure 35).
O
n-C5H11
O
n-C5H11
OH
n-C5H11
O
n-C5H11
OH
path A
path B
O
HO
n-C5H11
n-C5H11
OH
O
HO
n-C5H11
n-C5H11
OH
Rhodococcus sp. CBS 717.73
path Bpath A
95% ee
65% ee
53
Figure 35. Enzyme-catalyzed intramolecular 1,3-dipolar cycloaddition reaction.[135]
Another recent development reported by Faber et al. [136] is a biocatalytic
hydrogen-transfer reduction of halo ketones into enantiopure epoxides. The enzyme used
in this study is either Rhodococcus ruber as lyophilized cell catalyst or an alcohol
dehydrogenase prepared from Pseudomonas fluorescens DSM 50106 (PF-ADH).
2.3.2.6 Multienzymatic One Pot Reactions
The use of a multienzyme to catalyze organic reactions is an interesting approach
in the application of domino reactions. There is no limit to the number of enzymes that
can be used in a single reactor to produce a complex structure in a domino fashion. Since
1990, many studies have been reported on the use of multienzyme cocktails in the
synthesis of many natural products including the synthesis of β-D-glucuronides [137], 2’-
deoxy-N-acetyllactosamine [138], sialyl oligosaccharides [139], precorrin-5 [140],
N
OH
OEtO O
CO2EtOO
NO
O
CO2Et
H
O
NO
O
CO2Et
H
H
N
OH
O
+
+
Candida antarcticalipase
NEt3
52% yield93% ee
38% yield99% ee
54
sialylated antigen T-epitope [141], fluoroshikimic acids [142], cefazolin [143,144], and
aromatic D-amino acid [145].
Sheldon and his co-workers [146] reported a two step, one pot enzymatic
synthesis of cephalexin from D-phenylglycine nitrile in 2002. Two enzymes which are
nitrile hydratase and penicillin G acylase were used in this approach. First, the D-
phenylglycine was hydrated by nitrile hydratase to form the corresponding amide which
subsequently underwent acylation reaction with 7-aminodesaacetoxycephalosporanic acid
(7-ADCA) by penicillin G acylase to generate cephalexin (Figure 36).
Figure 36. Two enzymetic reactions for the synthesis of cephalexin.[146] Wong et al. [147] developed the four enzyme system for the synthesis of L-
fructose. In this study, L-glyceraldehyde was produced in situ from glycerol in the
CN
NH2 NH2
NH2
O
Nitrile HydrataseH2O
N
S
O
CO2H
H2N
7-ADCA
N
S
O
CO2H
HN
NH2
O
Penic
illin G
Acylas
e
Cephalexin
55
presence of galactose oxidase, catalase, rhamnulose-1-phosphate aldolase, and acid
phosphatase (Figure 37).
Figure 37. Four enzyme system for domino synthesis of L-fructose.[147]
Most recently, Kroutil et al. [148] reported the one pot, two step, two enzyme
cascade reaction for the synthesis of enantiopure epoxide. In this study, enantiopure (R)-
and (S)-epoxides were obtained by the reaction which combined either (R)- or (S)-
selective alcohol dehydrogenase with a non-selective halohydrin dehalogenase. First, the
pro-chiral α-chloro ketone was streoselectively reduced to the halohydrins as an
intermediate by alcohol dehydrogenase, and then the intermediate was converted to
epoxide by a non-enantioselective halohydrin dehalogenase (Figure 38).
Figure 38. Two enzyme system for the synthesis of enantiopure epoxide.[148]
OHHO
OHO
OH
OH OH
OH
OH
OH
OH
O
galactose oxidasecatalase
rhamnulose-1-phosphate aldolasethen acid phosphatase
H2O3PO OH
O
RCl
O alcohol dehydrogenaseNAD(P)H
OH OR
Cl
OH
*
(R) or (S)
O
R
(R) or (S)
halohydrin dehalogenase
- HCl
56
2.4 Laccase 2.4.1 Distribution in Nature Laccase (EC 1.10.3.2, p-diphenol:oxygen oxidoreductase) is an enzyme
belonging to the family of multicopper blue oxidase which typically found in plant and
fungi. Laccase can catalyze the oxidation of a variety of compounds including ortho and
para-diphenols, polyphenols, aminophenols, polyamines, lignins, aryldiamines, and a
number of inorganic ions, while reducing molecular dioxygen to water [12,149-152].
Laccase was first discover by Yoshida in 1883 in the sap of lacquer tree Rhus
vernicifera [153] and the enzyme has been characterized in great detail later in 2001 by
Huttermann et al. [154]. However, the report of laccase in other plant species is more
limited and partially characterized. These laccases include laccases form Rhus
succedanea [155], Acer pseudoplatanus [156], Pinus taeda [157,158], Populus
euramericana [159], Liriodendron tulipifera [160], Nicotiana tobacco [161], Lolium
perenne [162], and Zea mays [163]. In plant, laccase participates in the formation of
polymer lignin via radical-based mechanisms [156,164,165].
A few year later after the discovery of the plant laccase by Yoshida, fungal
laccases were discovered by Bertrand in 1896 [166]. The majority of laccases
characterized so far were isolated from fungi, and the reports of their presence in more
and more fungal species have been published [167,168]. Up to now, more than 100
laccases have been purified from fungi, and laccase from the wood-rotting white-rot
basidiomycetes were the most enzyme purified. The wood rotting fungi that produce
laccase are Trametes versicolor, T. hirsute (C. hirsutus), T. ochracea, T. villosa, T.
gallica, Cerrena maxima, Coriolopsis polyzona, Lentinus tigrinus, Pleurotus eryngii, etc.
57
Laccases have several roles in fungi including lignin degradation, morphogenesis, fungal
plant-pathogen/host interaction, and stress defence [8,167,168].
There are also some reports about laccase activity in bacteria [169,170].
Moreover, proteins with features typical of laccases have recently been identified in
insects [171].
2.4.2 Laccase Structure
Laccases are glycoproteins which often occur as isoenzymes that oligomerize to
form multimeric complexes. The molecular weight of the monomer ranges from about 50
to 130 kD. The carbohydrate moiety of laccases consisting of mannose, N-
acetylglucosamine, and galactose ranges from 10 to 45% of the protein mass in laccases.
This carbohydrate moiety is believed to be responsible for the stability of the enzyme
[3,152].
For the catalytic activity, the active site of laccases contains four copper atoms
which are one type-1 (T1) copper and a tree-nuclear cluster (T2/T3) consisting of one
type-2 (T2) and two type-3 (T3) coppers. T1 copper atom is located at the distance of
about 12 Å from the T2/T3 site, and T2 copper atom is located at the distance of about 4
Å from T3 copper atoms [172-174]. The T1 copper has a trigonal coordination with two
histidine and one cysteine, and the axial ligand of T1 is methionine in the bacterial
(CotA) [173] and leucine or phenylalanine in fungal laccases. The T1 copper confers the
typical blue color to multicopper proteins due to the strong absorption around 600 nm.
This intense absorption caused by the covalent copper-cysteine bond. Moreover, type-1
copper is the site where substrate oxidation takes place because of its high redox potential
58
of ca. +790 mV. Type-2 copper is coordinated by two histidines and type-3 coppers are
coordinated by six histidines. Type-2 copper shows only weak absorption in the visible
region and reveals paramagnetic properties in electron paramagnetic resonance (EPR)
studies. While type-3 coppers, a binuclear copper site with copper paired
antiferromagnetically through a hydroxyl bridge, exhibit the absence of an EPR signal.
The T3 site can be characterized by electron absorption at 330 nm (oxidized form)
[155,175,176]. In addition, the trinuclear cluster (T2/T3 site) is where the reduction of
molecular oxygen and release of water takes place. Figure 39 illustrated a scheme of
active site of laccase CotA from Bacillus subtilis.
Figure 39. Active site of laccase CotA from Bacillus subtilis (adapted from Enguita et al. [173]). Up to now, the three-dimensional structure [177] have been determined for five
fungal laccases from Coprinus cinereus (with the T2 copper removed) [178], Trametes
Met
Cys
His
His
His
His
His
His
His
His
His His
Cu1
Cu2 Cu3
Cu4
OH
HOH
Type-1 Copper
Type-3 Coppers
Type-2 Copper
59
versicolor [179,180], Pycnoporus cinnabarinus [181], Melanocarpus albomyces [182]
and Rigidoporus lignosus [174]. Moreover, the three-dimensional structure of laccase
CotA from endospores of Bacillus subtilis has also recently been published [173,183].
2.4.3 Catalytic Mechanism and Properties
Laccase catalysis is proposed to comprised three major steps [155,184,185]:
1. Type-1 copper is reduced by accepting electrons from the reducing substrate.
2. Electrons are transferred ~13 Å from type-1 copper to the trinuclear T2/T3
cluster.
3. Molecular oxygen is activated and reduced to water at the trinuclear T2/T3
cluster.
Figure 40 shows the catalytic cycle of laccase showing the mechanism of four-
electron reduction of a dioxygen molecule to water at the enzyme copper sites [186].
Dioxygen molecule interacts with the completely reduced trinuclear cluster (T2/T3) via a
2e- process (k ≈ 2 × 106 M-1s-1) to produce the peroxide intermediate which contains the
dioxygen anion [187]. One oxygen atom of the dioxygen anion bound with the T2 and T3
copper ions and the other oxygen atom coordinated with another copper ion of T3. Then,
the peroxide intermediate undergoes a second 2e- process (k > 305 s-1) [172], and the
peroxide O-O bond is splitted to produce a native intermediate which is a fully oxidized
form with the three copper centers in the trinuclear site mutually bridged by the product
of full O2 reduction with at least one Cu-Cu distance of 3.3 Å. This native intermediate
form of lacccase was confirmed by the combination of Cu K-edge x-ray spectroscopy
(XAS) and magnetic circular dichorism (MCD) studied by Solomon et al. [150].
60
Moreover, a combination of model studies and calculations has further demonstrated that
the three copper centers in the trinuclear cluster are all bridged by a μ3-oxo ligand [188].
This structure has a single μ3-oxo ligand bridging all three coppers at the center of the
cluster, with the second oxygen atom from O2 either remaining bound or dissociated from
the trinuclear site as shown in the native intermediate structure in Figure 40. This μ3-oxo
bridged structure of the native intermediate provides a relatively stable structure that
serves as the thermodynamic driving force for the 4e- process of O2 reduction, and also
provides efficient electron transfer (ET) pathways from T1 site to all of the copper
centers in the trinuclear cluster [188]. This efficient ET pathways lead to the fast
reduction of the fully oxidized trinuclear cluster in the native intermediate to generate the
fully reduced site in the reduce form for further turnover with O2. The native intermediate
can slowly convert to a completely oxidized form called “resting” laccase which has the
T2 copper isolated from the couple-binuclear T3 centers. The decay of the native
intermediate to the resting enzyme proceeds via successive proton-assisted steps as
illustrated in Figure 41 [189]. The first proton binds at μ3-oxo center and then the second
proton binds at T3 OH- bridge. Finally, the three copper centers in the trinuclear cluster
are uncoupled to form the resting form of laccase. The slow decay of the native
intermediate is due to the rearrangement of the μ3-oxo-bridge, the rate limiting step, from
inside to outside of the cluster. The T1 site of this resting laccase can be reduced by a
substrate. However, the electron-transfer rate onto the trinuclear cluster (T2/T3) is too
low to be significant for catalysis [150,155].
61
Figure 40. Catalytic cycle of laccase showing the mechanism of four-electron reduction of a dioxygen molecule to water at the enzyme copper sites (adapted form Shleev et al. and Solomon et al. [186,188]).
Cu1+
H2O
Cu1+ Cu1+
Cu1+
T2
T3
T1
Reduced Form
+ O2
Cu1+
H2O
Cu2+ Cu2+
Cu1+
T2
T3
T1
O
O
Peroxide Intermediate
Cu2+
OH
Cu2+ Cu2+
Cu2+
T2
T3
T1
O
OH
Cu2+
OH
Cu2+ Cu2+
Cu2*
T2
T3
T1
- H2O
+4e-
rapidly
slowly+4e-
Catalytic Cycle
Oxidized "Resting" Form
Native Intermediate
OH
O-O bond cleavage
(H)
62
Figure 41. Proposed decay mechanism of the native intermediate to the resting laccase.[189]
Laccase can catalyze the oxidation of a variety of compounds including ortho and
para-diphenols, polyphenols, aminophenols, polyamines, lignins, aryldiamines, and a
number of inorganic ions [12,149-152]. Laccase will abstract an electron from substrates
which produces a free radical, and reduce oxygen to water. The simplify scheme of
laccase-catalyzed redox cycles for substrate oxidation and the example of the oxidation of
hydroquinone by laccase are illustrated in Figure 42.
O
Cu2+
OH
Cu2+
Cu2+
OH(H)
T2
T3
Native Intermediate
H+
OH
Cu2+
OH
Cu2+
Cu2+
OH(H)
H+
O
Cu2+
O
Cu2+
Cu2+
OH(H)
HH
H
H2O
Cu2+
OH
Cu2+
Cu2+
OH
HO
H
HO
H
Resting Laccase
63
Figure 42. (a) Scheme of laccase-catalyzed redox cycles for substrate oxidation; (b) The example of the oxidation of hydroquinone by laccase. Fungal laccases typically exhibit pH optima in the range from 3.5 to 5.0 when the
substrates are hydrogen atom donor compounds, and the pH-dependence curve is bell-
shaped [190-197]. The optimum pH for phenolic compounds can actually increase at
higher pH to a limit. The limit for increasing the pH during substrate oxidation results
from the balance between the redox potential difference between the substrate and the
inhibition of the T2/T3 copper site by the binding of OH- ion [198,199]. The pH optimum
of plant laccases for substrates that are donors of hydrogen atoms was different from that
of fungal laccases. For example, laccase from Rhus vernicifera exhibited maximal
activity in neutral and weak alkaline solution [198].
The optimal temperature of laccases usually do not differ from other extracellular
ligninolytic enzymes with in the range from 50º to 70 ºC [168]. However, there are a few
Laccase(ox)
Laccase(red)
H2O
O2
Substrate(red)
Substrate(ox)
(a)
OH
OH
OH
O
4 4Laccase
O2
O
O
+
OH
OH
2H2O
non-enzymaticoxidization
2 2
(b)
dimerization orpoymeriation
Dimers or Oligomers or Polymers
Or
64
enzymes with the optima below 35 ºC such as the laccase from G. lucidum with the
highest activity at 25 ºC [200].
A wide spectrum of compounds has been described to inhibit laccase. These
inhibitors include small inorganic anions such as azide, cyanide, fluoride and hydroxide.
These ions bind with the T2/T3 site and this prevents the electron transfer from T1 site
onto T2/T3 site and inhibits the enzymatic activity [198,201]. Other inhibitors such as
metal ion (Hg+), fatty acids, quaternary ammonium detergents, have been shown to either
replace or chelate the copper centers, or de nature the protein [149].
2.4.4 Laccases in Organic Synthesis
Due to the catalytic and electrocatalytic properties of laccases, laccases have
received much attention from researcher in last decades as well as have shown the
potential of their wide application in several industrial and biotechnological processes
[21,152]. Moreover, laccases also pose the possibility of their application in fine organic
synthesis because of their ability to oxidize a variety of compounds [4]. The redox
potential of laccase is in the range of 0.5 to 0.8 mV (vs. normal hydrogen electrode
[NHE]) [198]. In the reactions where the substrate to be oxidized has a higher redox
potential than laccase or the substrate is too large to penetrate into the enzyme active site,
the presence of so-called ‘chemical mediator’ may be required to facilitate the reaction.
First, the mediator reacts with the laccase to form a strongly oxidizing intermediate.
Then, this oxidized mediator interacts with the bulky or high redox-potential substrate.
The mediators that are widely used such as N-hydroxybenzotriazole (HBT), 2,2’-
azinobis-(3-ethylbenzylthiozoline-6-sulphate) (ABTS), Violuric acid (VA), 3-
65
Hydroxyanthanilic acid (HAA), and 2,2,6,6-tetramethyl-1-piperidinyloxyl (TEMPO)
(Figure 43) [9,202]. However, this section will focus only on the laccase-catalyzed
reaction in the absence of mediators.
Figure 43. Chemical structure of laccase mediators.
2.4.4.1 Laccase-Catalyzed Oxidation Reaction
2.4.4.1.1 Laccase-Catalyzed Transformation of Phenolic and Other Compounds
Laccases have been reported to oxidize many phenolic compounds [198,203-207].
For example, Trejo-Hernandez and his co-workers [203] studied the use of laccase in the
crude extract of the residual compost of Agaricus bisporus to oxidize phenolic
compounds including guaiacol, 2,6-dimethoxyphenol, ventral alcohol, aniline, and
phenol. All tested substrates formed insoluble products after being oxidized except for
ventral alcohol that was transformed to a soluble aldehyde. The relative activity of the
compost extract was 2,6-dimethoxyphenol > guaiacol > phenol > ventril alcohol >
N
N
N
OH
HBT
HN
HN
O
O
O
N
OH
VA
N
OH
O
HAA
S
N
N N
N
S
O3S
SO3
ABTS
N
O
TEMPO
66
aniline. Recently, the product of the oxidation of 2,6-dimethoxyphenol by Rhus laccase
was determined for the first time by Wan et al. The reaction was conducted in water-
organic solvent system. They found that only one product, 3,3’,5,5’- tetramethoxy,1,1’-
biphenyl-4,4’-diol (Figure 44), was produced [206].
Figure 44. Stucture of 3,3’,5,5’- tetramethoxy,1,1’-biphenyl-4,4’-diol produced by laccase catalyzed the oxidation of 2,6-dimethoxyphenol.[206]
Monolignols including isoeugenol, coniferyl alcohol, and ferulic acid have also
been investigated for the laccase-catalyzed oxidation reactions. Chen and his co-workers
[208] studied the oxidation of isoeugenol and coniferyl alcohol by laccase from Rhus
vernicifera (tree) and Pycnoporus coccineus (fungus) in acetone-water (1:1, v/v). The
rate of Pycnoporus laccase-catalyazed oxidation of isoeugenol and coniferyl alcohol is
approximately 3 to 7 times faster than the rate of Rhus laccased-catalyzed oxidation. The
rate of the oxidation depends on the nature of both monolignol and laccase (Figure 45).
HO
MeO
MeO
OMe
OH
OMe
67
Figure 45. Dimer and tetramer products from the oxidation of isoeugenol alcohol by laccase.[208] Nishida and Fukuzumi [209] examined the transformation of ferulic acid by white
rot fungus, Trametes versicolor, in a medium containing glucose and ethanol under
aerobic condition. The ferulic acid was transformed into coniferyl alcohol,
coniferylaldehyde, dihydroconiferyl alcohol, vanillic acid, vanillyl alcohol, 2-
methoxyhydroquinone and 2-methoxyquinone. Falconnier et al. [210] also reported the
biotransformation of ferulic acid to vanillin by the white rot fungus Pycnoporus
cinnabarinus I-937 (Figure 46).
OH
OCH3
Isoeugenol
R. vernicif era laccase
acetone-water (1:1)23 oC, 24 h.
O
OCH3
OH
OCH3
+
OH
OCH3
CH
HC
CH3
O
H3CO
HO
OH
OCH3
CH
HC
CH3
O
H3CO
HO
+O
OCH3
OH
OCH3
O
H3CO
OH
H3CO
43% 11.7%
3.3%
1.7%
68
Figure 46. Biotransformation of ferulic acid by laccase.[210]
Figure 47. The synthesis of bis-lactone lignans.[211] The oxidation of ferulic acid by laccase was recently used to synthesize phenolic
colorants [212]. The oxidation was conducted in a biphasic hydro-organic system
consisting of ethyl acetate and sodium phosphate buffer to generate the intermediate
stable yellow products. This biphasic system facilitates the separation of the yellow
product which were soluble only in the organic phase and prevent the polymerization of
this intermediate to form brown polymer by reducing the activity of laccase in the
presence of organic solvent. They suggested that this yellow color compound can be used
OH
OCH3
COOH
Ferulic acid
COOH
OH
OCH3
+
CHO
OH
OCH3
Vanillic acid Vanillin
Laccase
(27.5%)
P. cinnabarinus I-937
6 days
R
OH
OMe
COOH
HO
MeO
R
O
O
H
H
O
O
R
OH
OMe
Sinapinic acid: R = OMeFerulic acid: R = H
Laccase, O2
EtOAc-buf fer
Bis-lactone lignansR= OMe (97%)R = H (36%)
69
as food colorants. However, this yellow compound is still in progress to elucidate the
structure. Moreover, the synthesis of bis-lactone lignan was reported to perform via the
transformation of sinapinic acid and ferulic acid by laccase in biphasic system (Figure
47) [211] .
Azo dyes, the largest group of colorants used in industry are able to oxidize by
laccase [213-215]. Renganathan and Chivukula [213] examined the oxidation of phenolic
azo dyes catalyzed by laccase from Pyricularia oryzae. Laccase oxidized azo dyes to 4-
sulfonylhydroperoxide, quinone compound, and other products (Figure 48). This study
suggests that laccase oxidation can result in the detoxification of azo dyes. Most recently,
Rehorek et. al. [214] reported a simultaneous combination of laccase and ultrasound
treatment in acetate buffer (pH 4.5) at 40 ºC for the degradation of azo dyes such as acid
oranges and direct blue dyes. The degradation process was monitor by UV-Vis
spectrometry and HPLC analysis. Compare to laccase or ultrasound treatment, the
stimultaneous treatment with laccase and ultrasound showed at least the same or higher
degradation rates of the azo dyes. Besides the degradation of azo dyes, laccase was also
reported to catalyze the formation of azo dyes by oxidative coupling between o-, m-, and
p-methoxyphenols and 3-methyl-2-benzothiazolinene hydrazone [216].
Figure 48. The oxidation of phenolic azo dyes by laccase.[213]
O3S N N
R1
OH
R2
R1 = CH3 or OCH3 or HR2 = CH3 or OCH3
O
O
R1 R2
+
OOH
SO3
4-Sulfophenylhydroperoxide
Laccase
Phosphate buffer pH 6.5
70
The transformation of other compounds such as steroid hormones [6,217,218],
alkaloids [219], flavonols [220], procyanidin B-2 [221], and N-(2-alkylamino-4-
phenylimidazol-1-yl)-acetamides [17] have been reported. The examples of these studies
are summarized in Table 3.
Table 3. Some examples of laccase mediated transformation of natural compounds.
Reaction Reference
[6] HO
OH
Steroid hormones β-estradiol
O
OH
HO
OH
+
O
OH
HO
HO
OH
HO
OH
+HO
OH
OH
HO
T. pubescens laccaseAcOEt-Acetate buffer pH4.5
rt, 48 h.
OH
14%
12.7%
71
Table 3. (Continued)
[219]
[220]
[221]
NMe
NEt
OAcOH
CO2Me
MeO
NMe
NEt
AcO
CO2Me
MeO O
NMe
NEt
AcO
CO2Me
MeO O
N NMe
OAc
O
CO2Me
OMe
+Laccase
Vindoline
OHO
OH O
OH
Galangin (flavonol)
T . versicolor laccase
OHO
OH OO
OHO
OH O
OH
OH
O
OH
O
OH
HO
OH
HO
OH
OH
OH
OH
OH
Procyanidin B-2
Laccasewater
O
OH
HO
OH
OH
OH
O OH
OH
OH
HO
O
Procyanidin A-2
72
Table 3. (Continued)
[17]
2.4.4.1.2 Lacccase-Catalyzed Oxidative Deprotection Reactions
Moreover, the use of laccase in oxidative deprotection for peptide synthesis has
been developed. A method to remove phenylhydrazide protecting group of both α- and γ-
carboxyl group by laccase have been proposed by Semenov et. al. [222]. The deblocking
method was performed under mild condition in aqueous medium and pH 7.0 in the
presence of oxygen. Therefore, this deprotection method lead to non-oxidative
modification without destruction of amino acid side chains. Recently, Rutjes and his co-
workers [223] reported the oxidative deprotection of p-methoxyphenyl (PMP)-protected
amines by laccase under mildly acidic condition (Figure 49). In addition, they found that
the use of mediators lead to an extension of the substrate scope and increase reaction rate.
N
N
NHPr
Ph
N-[4-phenyl-2-(propylamino)imidazole-1-yl]-acetamide
T . Versicolor laccase24 h
N
N
NHPr
Ph
HC
O
NHAc
NHAc
N
HN
NHPr
Ph
+
Bz NHN
N NHPr
NHAc
O Ph NH2
O
+
73
Figure 49. The oxidative deprotection of p-methoxyphenyl (PMP)-protected amines by laccase.[223] 2.4.4.1.3 Laccase-Catalyzed Oxidative Coupling for the Synthesis of the Pharmaceutical
Importance Compounds
Laccase have been reported to use for the synthesis of the pharmaceutical
importance compounds by oxidative coupling of the desired substrates to form the
corresponding dimer products. Some of the phenoxazinone chormophores having
antibiotic activity have successfully been synthesized via laccase-catalyzed oxidative
coupling reactions [224-227]. The synthesis of these phenoxazinone chormophores
involved the formation of aminophenoxy radicals by oxidation of o-aminophenols by
laccase at the first step. These radicals then underwent coupling and cyclocondensation
reaction to form the corresponding products. However, the reaction mechanism of this
synthesis is still under investigation. For example, actinocin, chormophore of
actinomycin antibiotics, was synthesized by laccase mediated oxidation of 4-methyl-3-
hydroxyanthranilic acid (4-M-3-HAA) (Figure 50) [224]. Laccase used in this study was
immobilized in polyacrylamide gel. The reaction proceeded successfully in aqueous
medium and in 60% acetonitrile.
HO
N
CH3
PMP
PMP = OMe
HO
NH2
CH3
+
O
O
LaccaseDMSO-Buffer pH 3.0
rt, 18 h
91% conversion
74
Figure 50. The synthesis of actinocin by laccase mediated oxidation of 4-methyl-3-hydroxyanthranilic acid.[224] Recently, Giurg et. al. [225] reported the synthesis of 2-amino-3H-phenoxazin-3-
one including actinocin, cinnabarinic acid, and questiomycin A by the catalytic oxidative
cycloaddition of o-aminophenols. These reactions were conducted in the presence of
laccase and oxygen in aqueous medium (Figure 51).
Figure 51. The synthesis of 2-amino-3H-phenoxazin-3-ones by the laccase catalyzed oxidative cycloaddition of o-aminophenols.[225]
COOH
NH2
OH
CH3
4-M-3-HAA
COOH
CH3
N
O O
COOH
NH2
CH3
immobilized laccase
0.1M phosphate buff er pH 5
Actinocin 74%
R1
NH2
OH
R2
R1
R2
N
O O
R1
NH2
R2
Laccase, Air
Water, pH 5.0, 20 oC
Questiomycin A: R1 = R2 = HCinnabarinic acid: R1 = COOH, R2 = HActinocin: R1 = COOH, R2 = CH3
0.5 - 23 h
75
The sulfonate analogue of cinnabarinic acid was recently synthesized by laccase
mediated the oxidative dimerization of 3-hydroxyorthanilic acid (Figure 52) [226].
Figure 52. The synthesis of the sulfonate analogue of cinnabarinic acid by laccase mediated the oxidative dimerization of 3-hydroxyorthanilic acid.[226] Forti and his co-workers [5] reported the transformation of trans-resveratrol
(3,5,4’-trihydroxystilbene) by laccase from Myceliophtora thermophyla and from
Trametes pubescens to generate the dehydrodimer product that has an antioxidant
properties (Figure 53).
Figure 53. The transformation of trans-resveratrol (3,5,4’-trihydroxystilbene) by laccase.[5]
SO3H
NH2
OH
SO3H
N
O O
SO3H
NH2
Laccase
water pH 6, 25 oC, 24 h
70%
HO
OH
OH
trans-resveratrol
O
HO
HO
OH
HO
OH
M. thermophyla laccase
dehydrodimer product
n-Butanol-Phosphate buffer pH 6.545 oC, 4 days
31%
76
Figure 54. The oxidation of a seires of hydroxystilbenes by laccase.[228]
R4
R5
R3
R2
R1
O R5
R2R3
R1
R4
R5
R3
R2
R1
1
23
4
5
6α
β
4
5
6 1
2
3
4-O-α-β-5 dimer
O
OH
R4
R5
R3
R2
R1
1
23
4
5
6α
β
R1
R2
R3
R5
4-O-β dimer
4
R4
R5
R3
R2
R1
1
23
4
5
6α
β
R1
R2
R3
4O
O 3
3-O-α-β-O-4 dimer
+
+
Laccase
EtOAc/ acetate buf ferpH 4.5, 40 oC
77
These authors recently reported the oxidation of a series of hydroxystilbenes,
analogues of the phytoalexin resveratrol by laccase from Trametes pubescens in ethyl
acetate/acetate buffer system [228]. In this study, three different dimeric product were
identified with the main product usually being 4-O-α-β-5 dimers. These products were
proposed to be generated via radical-radical coupling dimerization reactions (Figure 54).
Other biological active compounds have already been prepared. Antioxidant
gelatin-catechin conjugates have already been synthesized by the laccase-catalyzed
oxidation of catechin in the presence of gelatin in an aqueous medium [229]. Moreover,
the dimerization of Penicillin X [230], totarol [231], flavonolignan silybin [232], and
salicylic ester [13] by laccase have already been reported.
2.4.4.1.4 Laccase-Catalyzed Oxidative Cross-coupling Reactions
Laccases show to catalyze the oxidative cross-coupling reaction between different
molecules. Oxidative coupling of hydroquinone and mithramicine [233] or (+)-catechin
[16] have been examined. In the study of the cross coupling reaction between
hydroquinone and (+)-catechin, Rhus vernicifera laccase catalyzed the formation of two
new catechin-hydroquinone adducts (Figure 55). In this study, hydroquinone served as
both a shuttle oxidant and a reactant during laccase oxidations.
78
Figure 55. Laccase catalyzed the formation of catechin-hydroquinone adducts.[16]
O2
H2O
Laccase(red)
Laccase(ox)
OH
O
OH
OH
O
H
HOH
H
HO
OH
OH
OH
O
H
HOH
H
HO
OH
O
OH
O
HHO
+
(+)-Catechin
O
H
HOH
H
HO
OH
OH
OH
H
HO
OH O
H
HOH
H
HO
OH
OH
OH
H
OH
HO
6% 5.4%
79
Schauer et al. [15] reported the derivatization of the natural compound 3-(3,4-
dihydroxyphenyl)-propionic acid (dihydrocaffeic acid) via N-coupling reaction with
amines in the presence of laccase and oxygen in aqueous medium. The products of these
reactions were formed by a R-NH2 attack of a cation radical of dihydrocaffeic acid
(Figure 56). Later, they also studied laccase catalyzed a heteromolecular coupling of
dihydrocaffeic acid with 4-aminobenzoic acid in different reactor [234].
Figure 56. Laccase catalyzed N-coupling of dihydrocaffeic acid and amines.[15] A recent example of laccase catalyzed cross-coupling reaction is the synthesis of
Tinuvin, the benzotriazol base UV-absorber [235]. Laccase from Trametes hirsute was
used to catalyze the coupling reaction of 3-(3-tert-butyl-4-htdroxyphenyl)propionic acid
methylester to 1H-benzotriazole (Figure 57). This cross-coupling reaction occurred when
1H-benzotriazole was applied in four-fold molar excess.
OH
OH
OHO
NH2
O
HO
NH2
+
Or
Laccase, O2
Laccase, O2
HN
O
HO
OH
OH
OHO
NH
OH
OH
HO
O
Dihydrocaffeic acid
Acetate buffer pH 5
Acetate buffer pH 5
30 oC, 3 h
rt, 6 h
80%
60%
80
Figure 57. The synthesis of Tinuvin by a laccase-catalyzed reaction.[235]
Another recent example is the formation of protein-oligosaccharide conjugates
[236]. The formation of hetero-cross-coupling between tyrosine side chain of α-casein
and phenolic acid of hydrolyzed oat spelt xylan was catalyzed by laccase from Trametes
hirsula. This study shows another use of laccase in the modification of the biopolymer.
2.4.4.2 Laccase-Mediated Formation of Intermediate Quinones in Organic Synthesis
In this section, all reactions proceeded via the quinonoid intermediates of laccase
substrates. Laccase first oxidized the phenolic substrate to form phenolic radical which
further underwent nonenzymatic oxidation to generate quinonoid intermediate. The
quinonoid intermediate then reacted with other compounds to provide the corresponding
product (Figure 58).
OH
OO
NH
N
N
+T. hirsuta laccase
OH
OO
N
N
N
Tinuvin (5.1%)
Acetate buffer pH 4.51 h
81
Figure 58. Mechanism of laccase mediated the formation of quinonoid intermediate for Michael addition reaction.
Many studies of laccase-catalyzed synthesis of aminoquinones have been reported
[11,18,237]. Aminoquinones were synthesized by nuclear amination of p-hydroquinones
with primary aromatic amines in the presence of fungal laccase. The mechanism of these
reactions is likely to be proposed via Michael addition of primary amine to the quinoniod
intermediate (Figure 59a). In addition, this strategy also used to derivatize unprotected
amino acid L-tryptophane (Figure 59c) [238]. The laccase-catalyzed amination was also
used in the synthesis of bioactive compounds such as β-lactam antibiotic cephalosporins
(Figure 59d) [239] and novel penicellins (Figure 59e) [240]. Recently, Manda et al. [241]
showed that the quinonoid intermediate of laccase substrate can react with solvent such
as water, methanol, and other alcohols to form the C-O bond cross-coupling products
(Figure 59b). Besides laccase-catalyzed amination of p-hydroquinone, Laccase-catalyzed
amination of o-hydroquinone, such as laccase mediated Michael addition of 15N-
sulfapyridine to protocatechuic acid, have also been reported [242].
OH
OH
Laccase, O2
O
O
OH
O
2 2nonenzymatic
oxidation
OH
OH
+
O
O
Nu-H+Michael addition
O
O
Nu
82
Figure 59. Laccase mediated amination reaction.[11,238-241]
OH
OH
NH
OH
O
2Laccase
OH
O
NH
OH
O
2Nonenzymatic
Oxidation
O
O
NH
OH
O OH
OH
NH
OH
O
+
O
O
NH
OH
O
H2N COOH
O
O
R
NH
COOH
R=CONHCH2CH2OH
H2O or MeOH
O
O
NH
OH
O
OHor (OMe)
NH
OH
H2N
O
HN
OH
NH
O
O
O
R
R=CONHCH2CH2OH
NH2
O
HN
N
SH
O
COOH
CH3HO
Cefadoxil
O
O
NH
O
HN
N
SH
O
COOH
CH3HO
NH
OH
O
NH2
O
HN
N
H
OHO
S
COOH(H)
O
O
NH
OH
O
NH
O
HN
N
H
OHO
S
COOH(H)
+
+
+
+
+
(a)
(b)
(c)
(d)
(e)
Acetatebuffer pH5, rt70%
Laccase
Acetatebuffer pH5, rtLaccase
70%
Acetatebuffer pH5.6, rtLaccase
88%
Acetatebuffer pH5.6, rtLaccase
98%
Acetatebuffer pH5, rtLaccase
83
Laccase-mediated formation of intermediate quinone can be used in the domino
reaction. For example, Bhalerao et al. [243] reported laccase catalyzed one step synthesis
of 3-substituted-1,2,4-triazolo(4,3-β)(4,1,2)benzothiadiazine-8-ones (Figure 60).
Figure 60. The synthesis of 3-substituted-1,2,4-triazolo(4,3-β)(4,1,2)benzothiadiazine-8-ones by laccase mediated reaction of 5-substituted-4-amino-3-mercapto-1,2,4-triazoles and hydroquinone.[243] Recently, Leutbecher et al. [19] studied the synthesis of O-heterocycles via
laccase-catalyzed domino reaction between 4-hydroxy-6-methyl-2H-pyran-2-ones with
catechols. Moreover, Laccase initiated domino reaction of cyclohexane-1,3-diones with
catechols for the synthesis of 3,4-dihydro-7,8-dihydroxy-2H-dibenzofuran-1-ones has
been developed (Figure 61) [244]. The products yield ranging from 70% to 97%.
N
N
N
NH2
HS
R
+
OH
OH
Laccase
CH3CN-Phosphate Buffer pH 6.5(1:3)
N
NN
N
S
R
O
30 oC, 12 h
R = CH3, C2H5, Ph, Ph-OMe, Ph-CH3, Ph-Cl, Ph-Br
83-95%
84
Figure 61. Laccase initiated domino reaction of cyclohexane-1,3-diones with catechols.[244]
2.4.4.3 Laccase-Catalyzed Polymerization Reaction
Laccases have shown to catalyze polymerization reaction of many compounds
including acrylamide [245], 2-hydroxydibenzofuran [246], phenolic pollutants [247], 1-
naphtol [248,249], catechol [250], 4-cholroguaicol [251], Bisphenol A [252], and aniline
[253-255]. Some examples of these laccase catalyzed polymerization are shown in Table
4.
OH
OH
O
O
+Laccase, Air
pH 4-6, rt, 5h
O
O OH
OH
85
Table 4. Substrates, reaction conditions, and products from laccase catalyzed polymerization reactions.
Substrate Reaction condition Products Reference
Acrylamide
Laccase, water,
65 ºC, 4 h
Polyacrylamide (MW > 6 × 105) [245]
2-hydroxydibenzofuran
Laccase, acetate buffer pH 5, 30 ºC,
3 h
Dimers such as + Trimers and Oligomers
[246]
1-naphthol
Laccase, acetone-
acetate buffer pH 5, 25 ºC
Orange colored poly(1-naphtol)
Average MW = 4920 Da [248]
Bisphenol A
Laccase, phosphate
buffer pH 6, rt, 4 days
Dimer and Oligomers
[252]
O
NH2
O
OH O
OH
O
HO
+
O
O O
HO
OH
HO OH
HO OH
HO OH
86
Table 4. (Continued) Aniline + Sulfonated polystyrene (SPS)
Laccase, Citrate-
phosphate buffer
pH 3.5-4.4, 20 ºC
SPS-polyaniline complex [255]
In addition, many natural or artificial natural products have been synthesized by
laccase-catalyzed polymerization reaction. Kobayashi and his co-workers developed a
method for the preparation of artificial urushi [256-258]. Urushi is an insoluble polymeric
film formed by the crosslinking of urushiol monomer whose structure is a catechol
derivative with unsaturated hydrocarbon chain consisting of monoenes, dienes, and
trienes at 3-, or 4-position of catechol. The artificial urushi in this study was prepared by
laccase-catalyzed crosslinking of new urushiol analogues under mild conditions without
the use of organic solvents (Figure 62).
NH2
aniline
+
SO3 nSPS
87
Figure 62. The synthesis of artificial urushi by laccase-catalyzed polymerization of urushiol analogues.[258] Rutin is one of the most famous glycosides of flavanoid widely present in many
plants and has been reported to have biological activities including antioxidant,
antihypertensive, antiinflammatory, and antihemorrhagic activities. Therefore, Kobayashi
et al. [14] synthesized poly(rutin) by laccase-catalyzed oxidative polymerization of rutin
to amplify the antioxidant activity of rutin.
Figure 63. Structure of Rutin.
CH2O2CR
OH
OH
Laccase
O2
Urushiol Analogues
R = Or
"Artif icial Urushi"
O
OH
OH
O
O
HO
OH
OH
CH3
OH OH
O
O
OH
OH
OH
Rutin
88
These authors also synthesized poly(catechin) [259], a new class of flavonoid
polymers, via the polymerization of catechin by laccase in a mixture of acetone-acetate
buffer solvent. Poly(catechin) exhibited greatly amplified superoxide scavenging activity
and xanthine oxidase inhibitory activity compared with catechin. Moreover, Burton and
Ncanana [260] recently reported laccase-catalyzed polymerization of 8-hydroxyquinoline
to yield an antioxidant aromatic polymer (Figure 64). Eisenman et al. [261] reported the
use of Cryptococcus neoformans laccase to catalyzed the synthesis of melanin from both
D- and L-3,4-dihydroxyphenylalanine (DOPA).
Figure 64. The structure of poly(8-hydroxyquinoline).[260]
2.4.5 Laccase in Fiber Modification
Enzyme facilitated lignocellulosic fiber modification is recently a growing field of
research and interest [262]. Enzyme technology offers an environmentally friendly
method for modifying the fibers. Moreover, enzymatic treatment conditions are often
milder and less damaging to the fiber than chemical treatment. Laccase is one of the
enzymes used for the surface modification of lignocellulosic fibers [20,263,264].
N
OH
n = up to 15
Poly(8-hydroxyquinoline)
89
Fibers often have a set of their own properties. These properties, such as water-
binding capability, flexibility, rigidity, hydrophilicity, hydrophobicity, and the ability to
adhere to themselves and to other materials, depend on the structure and the composition
of the major components of the fiber which are cellulose, hemicellulose, and lignin [263].
Altering these fiber properties is a tremendous opportunity to produce a new value-added
material from this renewable resource.
The first part of this section will discuss the chemical composition and structure
of the lignocellulosic fibers. Next, the recent development in fiber modification by
laccase will be discussed.
2.4.5.1 Lignocellulosic Fibers
2.4.5.1.1 Chemical Composition
The three main natural polymers of lignocellulosic fibers are cellulose,
hemicellulose, and lignin.
Cellulose is a straight-chain polysaccharide composed of D-glucose repeating
units which are linked together by β-1,4-glycosidic linkages at the C1 and C4 positions as
shown in Figure 65 [265]. The degree of polymerization (DP) of cellulose in native wood
is around 10,000 but can decrease to less than 2000 after pulping [266]. The numerous
hydroxyl groups on the chain backbone of cellulose macromolecules lead to the
formation of both intermolecular and intramolecular hydrogen bonds. These hydrogen
bonds stiffen the straight chain and promote aggregation, forming a crystalline structure
[267]. Bundles of cellulose molecules are aggregated together in the form of microfibrils
90
with regions of high order (crystalline regions), and regions of low order (amorphous
regions). Microfibrils build up and form fibrils which form cellulose fibers.
Figure 65. Chemical structure of cellulose.[265] Cellulose has six crystalline polymorphs of which cellulose I and II are the most
commonly found [265,268]. Cellulose I, the native form produced in plant and other
organisms, is composed of parallel cellulose chains forming dense, highly hydrogen
bonded sheets. Natural cellulose I exists as two crystal phases, named Iα and Iβ. The
relative amount of Iα and Iβ depends on their origins. For example, some algae and
bacterial cellulose tend to be rich in Iα while cotton, wood, and ramie fiberstend to be rich
in Iβ [269,270]. Recently, Langan et al. [271,272] studied the crystal structure and
hydrogen-bonding system in cellulose Iα and Iβ from using synchrotron X-ray and
neutron fiber diffraction. They found that cellulose Iα and Iβ can both be described as
dense, highly hydrogen bonded sheets of parallel chains organized in sheet packed in a
“parallel-up” fashion. These two allomorphs show no hint of intersheet O-H···O hydrogen
bonding.The main difference between Iα and Iβ is the stacking of these sheets which is
displaced in the chain direction. The second sheet of both allomorphs is shifted in the
“up” direction by about c/4 relative to the first sheet. The third sheet in Iα is also shifted
O
H
O
H
HO
H
H
OHHO
OH H
O
H OH
O
HO
HO
H
H
O
H
H
HO
H
H
OHH
OHH
OH
O
HO
HO
H
H
H
n
91
up by about c/4 with respect to the second sheet, but in Iβ, it is shifted in “down” direction
by about c/4 relative to the second sheet. Therefore, there is a relative difference of about
c/2 in the position of the third sheet with respect to the second sheet in Iα and Iβ . These
authors also proposed that the most likely route for solid-state conversion of cellulose Iα
→ Iβ is the relative slippage by c/2 at the interface of the second and third sheets. They
also indicated that weak C-H···O hydrogen bonding also contributes to cellulose crystal
cohesion in both Iα and Iβ. There are more C-H···O inter-sheet bonds in Iβ than in Iα. This
contributes to the stability of Iβ over Iα.
Cellulose II consists of antiparallel cellulose chains that are arrange into less
dense sheets and shows to have hydrogen bonding both within sheets and between sheets
[273].
Hemicelluloses are branched heteropolysaccharides consisting of a number of
different sugar building units including glucose, xylose, mannose, galactose, and
arabinose (Figure 66). Hemicellulose is an amorphous polymer and this is attributed to
the low degree of polymerization (DP = 50-300), and the branch structure. Hemicellulose
is very hydrophilic, soluble in alkali, and easily hydrolyzed in acids [274]. The
proportions and the composition of hemicellulose vary from one species to another.
Hemicellulose content is typically 20-30% in softwood and 25-35% in hardwood [275].
Table 5 summarizes the DP and percentage of the major hemicelluloses in softwoods and
hardwoods. Galactoglucomannans and arabinoglucuronoxylan are the two main
hemicelluloses in softwood (Figure 67) while glucuronoxylan is the main hemicellulose
in hardwood [276].
92
Table 5. The degree of polymerization and percentage of the major hemicelluloses in softwoods and hardwoods.[277]
Hemicellulose type Percentage in wood (%)
Degree of polymerization
(DP) Galactoglucomannans 11-25 100 Softwoods Arabinoglucuronoxylan 7-10 100 Glucuronoxylan 15-30 200 Hardwoods Glucomannan 2-5 200
Figure 66. Sugar monomers in hemicellulose.
O
H
HO
H
HO
H
H
OHHOH
OH
β-D-Glucopyranose
O
H
HO
H
HO
OH
H
HHOH
OH
β-D-Mannopyranose
O
H
HO
H
HO
H
H
OHHOH
β-D-Xylopyranose
O
OH
H
H
HO
H
H
OHHOH
OH
β-D-galactopyranose
H
OH
H
HO H
H OH
O
HO
β-D-Arabinofuranose
93
Figure 67. Structure of hemicelluloses in softwood.[276]
Lignin, the second most abundant natural polymer on earth, is a complex aromatic
polymer most commonly derived from wood and an integral part of the cell walls of
plants. Lignin is totally amorphous and hydrophobic in nature. It gives rigidity to the
plants. Lignin macromolecule is a crosslinked three-dimentional phenolic polymer made
up of hydroxyphenylpropane units [278]. Due to the difficulty in isolating lignin without
modification, the original structure of native lignin is not yet known. However, numerous
information from lignin degradation products and model compound studies provides the
evidence that lignin formation originates from the polymerization of three different
hydroxyphenylpropane units known as monolignols. These monolignols are sinapyl,
coniferyl, and p-coumaryl alcohol as illustrated in Figure 68 [279].
OO
OH
HO
OH
OO
RO
OR
OO
OH
RO
OR
O
O
O
HO
OH
OH
OH
R = CH3CO or H
Galactoglucomannan
OO
HO
O
OO
HO OO
OO
OH
OH
H3COO
HO
HOOC
OH O
OH
OH
HOH2C Arabinoglucuronoxylan
94
Figure 68. The structure of monolignols.[279]
The polymerization of lignin is believed to proceed via the formation and
subsequent coupling of phenoxy radicals [278,280]. Figure 69 illustrates five main
resonance structures of the phenoxy radical which will undergo coupling reaction to form
a wide variety of linkages. The phenylpropane units are linked by C-C and C-O bonds.
Eight common interunit linkages in lignin are shown in Figure 70 [279]. Table 6 shows
the percentage of linkages found in hardwood and softwood lignin. The β-O-4 ether
linkage is the most abundant linkage in lignin, approximately 50% of total linkages in
softwood lignin. In addition, functional groups, including hydroxyl, methoxyl, and
carbonyl groups, have been identified in lignin.
CH2OH
R1
OH
R2
αβ
γ
Coniferyl alcohol (softwood/hardwood): R1 = OCH3, R2 = H
p -Coumaryl alcohol (softwood/hardwood): R1 = R2 = H
Sinapyl alcohol (hardwood): R1 = R2 = OCH3
95
Figure 69. Resonance structures of lignin precursors.[278]
Figure 70. Structure of eight different lignin linkages.[281-283]
CH2OH
OH
OCH3
-e, -H+
CH2OH
O
OCH3
CH2OH
O
OCH3
CH2OH
O
OCH3
CH2OH
O
OCH3
CH2OH
O
OCH3
OH
C
C
C
O
β-O-4
OH
C
C
C
α-O-4
O
OH
C
C
C
O
O
Dibenzodioxocin
OH
C
C
C
O
β-5
C
C
C
C
C
C
OH OH
5-5'
C
C
C
C
C
C
OH OH
β-β
C
C
C
OH
O
C
C
C
C
C
C
OH
OH
4-O-5 β-1
96
Table 6. The percentage of different lignin linkages in hardwood and softwood.[279,284]
Percentage found in wood (%) Type Name
Hardwood Softwood β-O-4 β-aryl ether 60 45 - 50 α-O-4 α-aryl ether 7 6 - 8 β-5 Phenylcoumaran 6 9 - 12 5-5’ Biphenyl and Dibenzodioxocins 7 18 - 25 4-O-5 Diphenyl ether 5 4 - 8 β-1 1,2-diphenylpropane 7 7 - 10 β-β β-β linked structures 3 3 2.4.5.1.2 The Effect of Kraft Pulping on Fiber Composition
The major chemical pulping process in North America is the kraft process. The
objective of any chemical pulping process is to remove enough lignin from cellulosic
fibers to produce a pulp suitable for the manufacture of paper and other related products.
In a conventional kraft cook, the wood chips are treated with an aqueous solution of
sodium hydroxide (NaOH) and sodium sulfide (Na2S), known as white liquor, in a large
pressure vessel called a digester. The white liquor and the wood chips are then heated to a
cooking temperature of about 170 ºC, typically reached after 1 – 1.5 hours. This allows
the cooking liquor to impregnate the chips. The cook is then maintained at the cooking
temperature for about 2 hours. Then, the contents are discharged into a blow tank to
disintergrate the softened chips into fibers [285]. During the kraft pulping treatment, the
hydroxide (OH-) and hydrosulfide anion (SH-), presenting in the pulping liquor, react
with the lignin. This reaction causes the lignin polymer to fragment into smaller
water/alkali-soluble fragment which are then dissolved as phenolate or carboxylate ions.
Hemicellulose and some cellulose are also chemically attacked and dissolve to some
97
extent. Typically, approximately 90% of lignin, 50% of the hemicellulose and 10% of
cellulose is removed in kraft pulping process [285].
The degradation of lignin during kraft pulping mainly proceeds through the
cleavage of ether linkages, with a concomitant generation of free phenolic hydroxyl
groups. The liberation of these phenolic hydroxyl group results in an increase of
hydrophilicity of the lignin and the lignin fragments. As a consequence, the solubility of
lignin in the pulping liquor is increased. However, the carbon-carbon linkages are more
stable and tend to remain after the pulping process. At the end of kraft pulping, the
remaining or residual lignin content is typically about 4-5% (by weight) [280,285].
Chakar and Ragauskas [280] recently reviewed the softwood kraft lignin process
chemistry. Two main lignin reactions, which are degradation and condensation reactions,
occur during kraft pulping. The major degradation reactions are the cleavage of α-aryl
and β-aryl ether bonds [286]. α-Aryl lingkages are shown in Figure 71. The quinone
methide intermediate is formed after the α-aryl bond cleavage. This quinone methide
intermediate can react with SH- to generate a benzyl mercaptide structure. Then, the
mercaptide anion attacks the β-carbon to yield a thiirane intermediate and eliminates the
β-aryloxy group as illustrated in Figure 71. In addition, the terminal hydroxymethyl
group of the quinone methide intermediate can be eliminated as formaldehyde to yield an
alkali-stable enol ether (Figure 71) [287,288]. The cleavage of the β-aryl ether bond is
summarized in Figure 72. This cleavage involves the attack of an ionized hydroxyl group
present on the α- or γ-crabon.
98
Figure 71. Alkaline cleavage of α-aryl ether bond, sulfidolytic cleavage of β-aryl ether bonds in phenolic arylpropane units, and conversion into enol-ether units of quinone methide intermediates.[280]
MeO
O
HC
HC O
CH2
HO
OR
MeO
R = H, Aryl
-OR
α-aryl ether bond cleavage
MeO
O
CH
HC O
CH2
HO MeO
Quinone methide intermediate
-CH2O-H+
MeO
O
CH
HC O
MeO
Enol ether
+HS-
-H+
MeO
O
CH
HC O
CH2
HO MeO
-S
O
MeO
-
MeO
O
CH
CH
CH2
HO
S-S
MeO
O
CH
CH
CH2
HO
99
Figure 72. β-aryl ether bond cleavage in nonphenolic arylpropane unit.[280]
During the kraft pulping, the quinone methide intermediate acts as an acceptor
which can react with necleophiles such as SH-, OH-, and lignin nucleophiles (e.g.,
carbanions from phenolic structures). Therefore, these nucleophiles compete for quinone
methide intermediates. The condensation reaction proceeds via Michael addition between
quinone methide intermediate and phenolated ion, followed by the abstraction of a proton
and rearomatization to form the corresponding product. However, when the structures
contain a good leaving group, such as an aroxyl group, at the β-carbon, the cleavage of β-
aryl ether linkages will predominate over condensation reactions [280]. Figure 73
summarizes the proposed competitive addition of these necleophiles.
MeO
O
CH
HC O
CH2
O MeO
β-aryl ether bond cleavage
O
O
MeO
-
MeO
O
CH
CH
CH2
O
O
OH-
MeO
O
CH
CH
CH2
O
O
OH
100
Figure 73. Competitive addition of external (SH-) and internal (phenolate ion) nucleophiles to quinone methide intermediates.[280] Moreover, the carboxylic acid group content of the residual lignin is affected by
the kraft pulping process. Froass, Ragauskas, and Jiang [289] reported that the carboxylic
acid group content of the lignin increases as delignification proceeds. The enhancement
of carboxylic groups in residual lignin after kraft pulping is also reported by Jiang and
Argyropoulos [290]. This enhancement is accompanied by a decrease in the amount of
aliphatic hydroxyl groups.
Polysaccharides, including hemicellulose and cellulose, are also degraded during
the kraft process. The hemicellulose content is reduced by approximately 40%. The
dissolution of hemicellulose is caused by the combination of peeling and alkaline
hydrolysis reactions. The peeling reaction can be ended via the stopping reaction which
MeO
O
CH
HC R
+SH-
-H+
MeO
O
HC
HC R
S-
MeO
O
HC
HCS
-R-
R = aryloxy group
O
OMe
MeO
O
HC
HC R
O
H
OMe
-H+
MeO
O
HC
HC R
O OMe
R = aryl group
101
converts the reducing end group to a stable carboxylic acid group [276]. Therefore, it can
be assumed that virtually all carbohydrate end groups have been converted to carboxylic
acids at the end of kraft pulping. Figure 74 summarized the peeling and stopping
reactions of polysaccharides during kraft pulping.
10% of cellulose is removed during the kraft pulping process. This low loss of
cellulose is due to the low accesssability of OH- into the crystalline region of the
cellulose. In addition, about 90% of the extractives in wood are removed [285]. Table 7
shows yield values for individual wood composition after kraft pulping of Scots pine
(Pinus sylvestris, softwood) and birch (Betula verrucosa, hardwood).
102
Figure 74. Scheme illustrates peeling and stopping reactions of polysaccharides during kraft pulping. [291]
CH O
H C
CH
H C
H C
O H
HO
O R
O H
R '
CH O
C
CH
H C
H C
O H
O R
O H
R '
CH O
C
CH 2
H C
H C
O
O R
O H
R '
CO O H
H C
CH 2
H C
H C
O H
O R
O H
R '
CH 2O H
C
CH
H C
H C
O
HO
O R
O H
R '
CH 2O H
H C
C
H C
H C
O H
O
O R
O H
R '
CH 2O H
H C
C
H 2 C
O H
O
O R
CH O
R '-
CH 2O H
C
C
H 2 C
O
O
O R
CO O H
C
H 2 C
O H
C H 3
O R
-RO H
CH 2O H
C
C
CH
H C
O
HO
O H
R '
CH 2O H
C
CH 2O H
O +
CH O
H C O H
R '
CH O
H C
CH 2O H
O H
CH O
C
C H 2
O H
CH O
C
CH 3
O
CO O H
H C
CH 3
O H
CH 2O H
C
C
CH 2
H C
O
O
O H
R '
CO O H
C
CH 2
H C O H
R '
O H
C H 2 O H
CH O
H C
C
CH 2
H C
O H
O
O H
R '
-HC OOH
CH 2O H
C
CH 2
H C
O
O H
R '
CH 2O H
C
CH
CH
O
R '
CH O
C
CH 2
CH 2
O
R '
CO O H
CH O H
CH 2
CH 2
R '
R = Po lysa ccha rid e chainR ' = C H 2O H f or ce llulo se or g luco m an nan o r H f or x ylan
103
Table 7. Yield values for individual pulp components after kraft pulping of Scots pine (a softwood) and birch (a hardwood).[292]
Yield (%, on dry-wood basis)
Pine Birch Wood Component
Original After pulping Original After pulping Cellulose 39 35 40 34 Glucomannan 17 4 3 1 Xylan 8 5 30 16 Other carbohydrates and various components 5 - 4 -
Sum of carbohydrates 67 44 74 51 Lignin 27 3 20 2 Pitch 4 0.5 3 0.5 Sum of components (yield) 100 47 100 53
2.4.5.1.3 Structure of Lignocelluosic Fibers
Lignocellulosic fibers are composed of hollow cellulose fibrils held together by a
lignin and hemicellulose matrix. The cell wall of a fiber has a complex, layered structure
as illustrated in Figure 75. The hollow center of the fiber called lumen, and the sublayers
of the cell wall consisting of a thin primary wall and a thicker secondary wall. The
primary wall has a lower amount of cellulose and a higher amount of lignin compared to
the secondary wall. Cellulose microfibrils from the primary wall are organized in a loose
network almost perpendicular to the cell axis. The secondary wall is made up of three
layers, S1, S2, and S3 [293]. The secondary wall’s microfibrils have a parallel
arrangement. Each layer of the secondary wall has a different microfibrillar angle, the
angle between the fiber axis and the microfibrils. The microfibrillar angle in S1, S2, and
S3 layers are 50-70º, 10-30º, and 60-90º, respectively [293]. The microfibrils, providing
mechanical strength to the fiber, are made up of 30-100 cellulose molecules in extended
104
chain conformation. The thick S2 layer determines the mechanical properties of the fiber.
The amorphous phase in the cell wall consists of hemicellulose, lignin, and in some cases
pectin. The hemicellulose molecules are bonded with cellulose microfibrils by hydrogen
bonding. This cellulose-hemicellulose network is believed to be the main structure
component of the fiber cell. The compound that binds the two adjacent primary walls
together is called the middle lamella. The middle lamella (ML) is primarily composed of
lignin that holds the fibers together in the wood ultrastructure. The length of typical
softwood fibers is approximately 2.5-7.0 mm and the width is approximately 25-50 μm.
Typical hardwood fibers are approximately 0.8-1.6 mm long and 14-40 μm wide.
Figure 75. A softwood tracheid (fiber) cell wall structure (Adapted from Coté [294]).
105
2.4.5.2 Laccase Application in Fiber Modification
Recently, laccase research studies have shifted toward fiber modification.
Lignocellulosic fibers compose of lignin, the macro phenoxylic structure which can be
oxidized by laccase to form the phenoxylic radical in the fibers. These radicals appear to
undergo polymerization with each other or undergo coupling reaction with other
compounds. Therefore, they have been used to graft a variety of substrates onto the fiber
which leads to the alteration of fiber surface. Moreover, depending on the grafting
materials, the properties of the modified fibers can be designed to suit the end product.
Laccases have been applied for bonding of fiberborads, particle boards, paper
boards, and kraft-liner board [295-298]. The auto adhesion of wood fiber and particles
has been achieved using laccase for activation of the surface lignin. Laccase first oxidized
lignin at the surface fibers to generate the lignin phenoxy radicals. These radicals then
underwent the crosslinking reaction to form a crosslinked-network of lignin between
fibers. Laccase-catalyzed polymerization of lignin through cross-linking of lignin
phenoxy radicals led to the bonding and strength enhancement of lignocellulosic
materials. Recently, the internal bonding of particle boards was improved by laccase-
catalyzed funtionalization with 4-hydroxy-3-methoxybenzylurea [299]. In this study, 4-
hydroxy-3-methoxybenzylurea was used as a functional compound to graft with spruce
wood particle by laccase. The presence of the urea group in this funtionalized wood
particle led to crosslinking between the funtionalized wood particles and resin in
subsequent glueing processes (Figure 76), which improved the strength properties of the
particle boards.
106
Figure 76. Laccase catalyzed grafting of lignin with 4-hydroxy-3-methoxybenzylurea, followed by chemical crosslinking to urea/formaldehyde (UF) resin in the subsequent glueing process.[299]
Besides catalyzing auto cross-linking between lignin, laccases have been used to
catalyze the grafting reaction of various materials onto technical lignin. For example,
guaiacol sulfonate has been grafted onto lignin by laccase resulting in an increase of the
water solubility of lignin [22]. This reaction was initiated by an oxidation of lignin and
guaiacol sulfonate by using laccase to generate phenoxy radicals of both components.
These radicals then underwent the coupling reaction with each other to form guaiacol
sulfonate-grafted lignin. Huttermann et al. reported that the lignin phenoxy radicals
HN NH2
O
OCH3
OH
+ Wood ligninLaccase
pH 5, 25 oC90%
HN NH2
O
OCH3
OH
Wood lignin
U/F resin
N N
O
OCH3
OH
Wood lignin
N
O N N
N
O
N
O
107
formed by the laccase catalyzed oxidation reaction are so active that they can also react
with nucleophlies such as cellulose and starch. Therefore, this study shows that
carbohydrate can be covalently bonded with lignin via the laccase catalyzed reaction of
lignin with cellulose [23]. Moreover, Mai et al. reported many studies involving the
grafting of lignin with synthetic polymers derived from acrylic and acrylamide to create a
new class of engineering plastics [24-27]. The presence of both laccase and peroxides
such as dioxane peroxides were essential in the copolymerization of acrylamide and
acrylic with lignin. In addition, the results from many experiments, such as solubility
testing, elemental analysis, UV-Vis, FT-IR, and 13C-CPMAS spectroscopy, provided
evidence of grafting. In case of acrylamide-lignin copolymer, when freeze-dried this
copolymer appeared as homogeneous fibril-like particulates. The proposed mechanism of
the enzymatical grafting is illustrated in Figure 77.
108
Figure 77. Proposed mechanism of chemoenymatically induced graft copolymerization between lignin and acrylamide. [25]
O H
LigninLaccase
O
Lignin
O
Lignin . . . (1)
ROOH + e- RO + OH-
Phenoxy Quinone
a)
b) ROOH
Phenoxy Phenol
ROO + H+ + e-
(2)
RO(O) + (n+1)CH2=CHCONH2 RO(O)(CH2CH(CONH2))nCH2CH(CONH2) (3)
O
Lignin RO(O)(CH2CH(CONH2))nCH2CH(CONH2)+
O
Lignin
(CH(CONH2)CH2)n+1(O)OR
O H
Lignin
(CH(CONH2)CH2)n+1(O)OR
(4)
109
In addition, lignocellulosic fibers have been reported to be grafted with a variety
of low molecular weight compounds. Chandra et al. modified high-lignin softwood kraft
pulp by grafting with phenolic acids (Figure 78), including 4-hydroxyphenylacetic acid
(PAA) [30], 4-hydroxybenzoic acid (4-HBA) [31], and gallic acid [29], in the presence of
laccase. The grafting of these phenolic acids was performed in water (pH 4.5) at 45 ºC for
2-4 hours and resulted in an increase of carboxylic acid groups, water retention, tensile
strength, and burst strength of the resulting paper. Table 8 summarizes some of the paper
strength test results of the phenolic-grafted pulp experiments. The strength increases were
due to the improvement of hydrogen bonding between fibers and the cross-linking
between phenoxy radicals within the sheet.
Figure 78. Phenolic acids for the modification of high kappa pulp.
OH
COOH
PAA
COOH
OH
4-HBA
COOH
HO
OH
OH
Gallic acid
110
Table 8. Paper strength test result for high lignin kraft pulp treated with laccase and phenolic acids.
Treatment
Physical properties of paper Control Laccase
Phenolic acid
Laccase + Phenolic
acid 4-Hydroxybenzoic acid (4-HBA)-treated experiment [31]
Apparent density (g/cm3) 0.43 0.44 0.42 0.47 Burst index (kPa.m2/g) 2.38 2.39 2.42 2.95 Tensile index (N.m/g) 36.65 38.87 36.98 42.10
4-Hydroxyphenylacetic acid (PAA)-treated experiment [30] Apparent density (g/cm3) 0.38 0.39 0.38 0.39 Burst index (kPa.m2/g) 1.76 2.10 1.76 2.16 Tensile index (N.m/g) 31.40 33.46 30.56 34.54
Gallic acid-treated experiment [29] Apparent density (g/cm3) 0.41 0.42 0.42 0.43 Burst index (kPa.m2/g) 2.46 2.40 2.41 2.68 Tensile index (N.m/g) 33.9 33.8 34.0 40.3 Wet tensile index (N.m/g) 1.38 1.74 1.21 2.26 Viikari et al. [28] reported the modification of the fiber surfaces of
thermomechanical pulp (TMP) by laccase and tyramine via a two-stage functionalization
method. This method consists of an enzymatic activation of fiber surfaces followed by
the addition of radicalized compounds that react preferentially through radical coupling.
The degree of bonding in this study was determined by electron spectroscopy for
chemical analysis (ESCA) which showed an increase in nitrogen content which
originated from nitrogen in tyramine. The results showed that the nitrogen content of
laccase-tyramine treated unbleached and bleached TMP increased to 0.6% and 1.5%,
respectively. In addition, the FTIR spectra of tyramine-grafted samples indicated the
111
formation of ether linkages at 1060 cm-1. Therefore, the authors suggest that tyramine
was bond by ether linkage to the pulp. The proposed structure of the modified fiber is
illustrated in Figure 79. The mechanism was suggested to start with one electron
oxidation of the phenolic hydroxyl groups of both lignin and tyramine to generate the
corresponding radicals. These radicals then react via a radical coupling reaction to form
the corresponding tyramine-bonded lignin (Figure 80).
Figure 79. The proposed structure of the modified TMP with tyramine by laccase.[28]
O
Lignin
OH
OCH3
H3NH2C
112
Figure 80. Proposed mechanism for grafting of tyramine to lignin by laccase.[28]
Recently, Elegir and his co-workers developed antimicrobial cellulose packaging
through laccase-mediated grafting of antimicrobial active phenolic compounds, such as
caffeic acid and isoeugenol, with unbleached kraft liner fibers [300]. Schroder et al. [301]
reported the grafting of lignocellulosic surfaces with methoxyphenols and hydroquinone
catalyzed by laccase to generated color and bacterial resistant lignocellulosic fibers.
Moreover, Kim et al. [302] examined enzymatic polymerization on the surface of
functionalized cellulose fibers. In Kim’s study, laccase catalyzed the polymerization of
catechol on the surface of aminized cellulose to from polycatechol-coated aminized
cellulose (Figure 81).
Lignin
OMe
OH
OH
OH
CH2NH3+Cl-
Tyramine
Laccase, O2
Lignin
OMe
O
Lignin
OMe
O
. . . +
O
OH
CH2NH3+Cl-
Radical coupling
Lignin
OMe
O
O
-Cl+H3NH2C
OH
Lignin
OMe
OH
O
-Cl+H3NH2C
OH
113
Lignin itself has also been reported to be grafted onto lignocellulosic fibers. For
example, an ultra-filtered lignin isolated from kraft black liquor was linked with kraft
liner pulp and chemi-thermo-mechanical pulp by laccase from Trametes pubescens. This
modification provided more than a twofold increase in wet strength of kraft liner pulp
[303].
Figure 81. Laccase catalyzed Coupling reaction of aminized cellulose with catechol.[302]
O
O
O
O
OH OH OH OH
OHO
CH2CH2 S
O
O
NH2
n
Laccase, O2, Catechol
O
O
O
O
OH OH OH OH
OHO
CH2CH2 S
O
O
HN
n
O
OH OH n
Aminized Cellulose
Coated Cellulose
114
2.4.6 Conclusions
Due to their high stability, selectivity for phenolic substructures, and mild
reaction conditions used in laccase-catalyzed reactions, laccases are attractive for fine
chemical synthesis and numerous synthetic processes have now been reported. A number
of the laccase catalyzed reactions provide routes for the synthesis of biologically active
compounds that have pharmaceutical significance. Moreover, the use of laccase as a
biocatalyst in the synthetic methods is primarily used to develop more environmentally
friendly processes when compared to the usual chemical-based synthetic processes that
involve the use or disposal of harzardous chemicals. The laccase catalytic processes
produce water as the sole-by product, and therefore could be ecologically friendlier. For
example, the chemical synthesis of phenoxazine derivatives involves the condensation of
the highly toxic, nitroso compounds, at elevated temperatures. Therefore, laccase was
used instead of chemical reagent to catalyze the synthesis of phenoxazines in water at
ambient temperature to provide greener synthetic method [225,226].
The laccase-catalyzed reactions are comparable to the chemical routes regarding
to reaction rate, purity of the products, stability of the products in the reaction medium,
and yields. For example, the formation of products from the nuclear amination reaction-
catalyzed by laccase is comparable with reaction using sodium iodate as oxidant [237].
However, there are still some disadvantages of using laccase in the organic synthesis
including the presence of buffer salts and protein in reaction medium makes the isolation
process more difficult, the price of laccase is more expensive than chemical reagents, and
the requirement of sufficient amount of oxygen for the catalytic system.
115
Apart from the use of laccase in organic synthesis, laccase-assisted modification
of wood also has potential in the area of the forest products industry. The main benefits
of laccase-catalyzed biografting of molecules to wood fibers are probably the absence of
harmful solvents and chemicals and the mild reaction conditions. Due to the versatility,
non-toxicity, and mild application conditions of laccase technology, laccase is likely to
remain the subject of intensive investigations in many areas of biocatalyst applications.
2.5 Lipases
2.5.1 A General Account
Lipases (EC 3.1.1.3, triacylglycerol hydrolase) belong to the family of hydrolases
that act on carboxylic ester bonds. Their physiological role is to catalyze the hydrolysis of
triglycerides to diglycerides, monoglycerides, fatty acids, and glycerols. They can also
catalyze the formation of acylglycerols from free fatty acids and glycerol (Figure 82)
[304-306].
Figure 82. Lipase-catalyzed reactions of triacylglycerols.[307]
CH2OCOR1
CHOCOR2
CH2OCOR3
+ 3H2Olipase
CH2OH
CHOH
CH2OH
+R1COOHR2COOHR3COOH
+ 2R4COOH
CH2OCOR1
CHOCOR2
CH2OCOR3
lipaseCH2OCOR4
CHOCOR2
CH2OCOR4
R1COOHR3COOH
+
116
Lipases are widely found in animals, plants, and microorganisms [308,309].
Currently, several lipases are commercially available. The majority of commercial lipases
are produced by fungi, yeast, and bacteria because of the ease of cultivating these
microorganisms on a large scale. In general, lipases are extracellular-acidic
glycoproteins. The molecular size of lipases is between 20 and 60 kDa [304]. Structural
characteristic include an α/β-hydrolase fold and a nucleophilic elbow where the catalytic
serine is located [307,310]. In addition, most lipases contain a ‘lid’ which is a helical
oligopeptide that shields the active site. This lid will open to provide free access for the
substrate when the enzyme interacts with a hydrophobic interface such as a lipid droplet.
Therefore, lipase changes into an activated form by substrate activation at the lipid-water
interface. This phenomenon is called interfacial activation and is unique structural
characteristic of this class of enzymes [304,311].
Lipases can be classified into three major groups according to their ability to
hydrolyze glycerides [304]. The first group is termed as 1,3-specific because they can
hydrolyze only the terminal positions of triglycerides. Since their substrate range is not
limited to triglycerides, this group can be regarded as lipases capable of hydrolyzing
primary and to a small extent secondary esters. Lipases in this group include lipases of
Rhizopus and Rhizomucor. The second lipase group can be termed as nonspecific because
they can hydrolyze both primary and secondary esters. The last group consists of those
few lipases that are positionally nonspecific but show fatty acid selectivity, cleaving only
ester bonds wherein the fatty acid is of particular type. In addition, lipases may also
exhibit chain length specificity.
117
In general, most animal lipases exhibit pH optima on the alkaline side, pH 8.0 -
9.0, while most microbial lipases show maximum stability in the neutral pH range [312].
Most lipases are optimally active at temperatures between 30 and 40 ºC [304]. Usually,
animal and plants lipases are less thermostable than the microbial extracellular lipases
[313].
The broad synthetic potential of lipases is largely because they possess broad
substrate specificity and tolerate organic solvents. Substrates other than triglycerides
include aliphatic, alicyclic, bicylic, and aromatic esters. Moreover, a wide range of
thioesters and activated amines can also be substrates for lipases. Lipases can be
employed for a variety of reactions such as esterification, interesterification, acidolysis,
alcoholysis, and aminolysis (Figure 83) [304,307,311,314-317]. In addition, lipases do
not require cofactors, and usually exhibit high chemoselectivity, regioselectivity, and
enantioselectivity. These properties make lipases the most versatile biocatalyst. Besides
the application of lipases in synthetic chemistry, the application of lipases are also found
in the detergent, food, leather, textile, oil and fat, cosmetic, paper and pharmaceutical
industries [305,318,319].
118
Figure 83. Examples of lipase-catalyzed reactions.[304]
H yd rolysis
R' O R ''
O
+ H 2Ol ipase
R ' O H
O
R ''O H+
E ste rif ic atio n
R' O H
O
R ''O H+ l ipaseR ' O R ''
O
+ H 2O
T ran sest erif icat ion
A cid o lysi s
R' O R ''
O
R ' '' O H
O
+ l ipase
R ''' O R ''
O
R ' O H
O
+
A lco ho lys is
R' O R ''
O
+ l ipase
R' O R '' '
O
+R '' 'O H R ''O H
I nte resterif i cati on
R' O R ''
O
R ' '' O R '' ''
O
+ l ipase
R' '' O R ''
O
R ' O R '' ''
O
+
A m inoly sis
R' O R ''
O
+ l ipase
R' N H R '' '
O
+R '' 'N H 2 R ''O H
119
Besides catalyzing the reactions in Figure 83 above, lipases are reported to
catalyze the Michael addition reaction, the 1,4-addition of a nucleophile to an α,β-
unsaturated carbonyl compound. The proposed mechanism reported to involve the
stabilization of the negative charge of the transition states in the oxyanion hole of the
active site, and the His-Asp pair serves as a proton shutter. The following section will
focus on the Michael reaction catalyzed by lipases.
2.5.2 Lipase-Catalyzed Michael Reaction
In 1986, Kitazume et al. [320] showed the possibility of hydrolases including
lipase from Candida cylindracea to catalyze Michael addition reactions. In this study,
optically active aliphatic and heterocyclic compounds possessing a trifluoromethyl group
were synthesized via an enzymatic chiral Michael addition reaction of 2-
(fluoromethyl)propenoic acid. The reactions were conducted in buffer solution pH 8.0
(Na2HPO4 and KH2PO4 solution) at 40 ºC and yielded the chiral products in the range of
40 to 90 % (Figure 84).
120
Figure 84. Asymmetric Michael addition reaction of 2-(trifluoromethyl)propenoic acid catalyzed by lipase from Candida cylindracea (* represents chiral center).[320] Torre et al. [321] has reported that lipase B from Candida antractica (CAL-B) can
catalyze a Michael addition of a secondary cyclic and non-cyclic amine to acrylonitrile.
The reactions were conducted in toluene at 30 ºC. In the presence of CAL-B, the rate of
the reactions were up to 100-fold faster than the reaction in absence of the biocatalyst.
The proposed mechanism of this process is summarized in Figure 85. The mechanism
starts with the accommodation of acrylonitrile in the active site. Then, the conjugated
addition of the necleophile leads to a zwiterionic intermediate stabilized by both the
oxyanion hole and the His-Asp pair. This His-Asp pair catayzes proton transference from
the incoming nucleophile to the α-carbon. Finally, a new acrylonitrile molecule shifts the
final product, leading to a new catalytic cycle.
CO2H
CF3
+ Nu-H NuCO2H
CF3
*Lipase (Candida cylindracea)
buffer pH 8.0, 40-41 oC
Nu-H Time (h) Yield (%) % e.e.
H2O 34 48 70Et2NH 92 47 71PhNH2 40 76 39
CO2H
CF3
+Lipase (Candida cylindracea)
buffer pH 8.0, 40-41 oC
NH2
YH NH
Y
O
CF3*
Y Time (h) Yield (%) % e.e.
O 21 83 41S 20 86 47N 24 56 38
121
Figure 85. Proposed mechanism of lipase catalyzed Michael addition of pyrrolidine and acrylonitrile.[321]
NH +CN
CAL-BToluene, 30 oC
NCN
H H H
N
C
Oxyanion hole
N
N
His
H O
O
Asp
NH
H H H
N
C
N
N
His
H O
O
Asp
NH
H H H
N
C
N
N
His
H O
O
Asp
N H
H H H
N
C
N
N
His
H O
O
Asp
N
CN
NCN
122
Recently, Dai and his co-workers [322] showed the ability of lipase M from
Mucor javanicus in the synthesis of pyrimidine derivatives containing a branched sugar
which may possess potential antitumor and antivirus activities. In this study, lipase M
catalyzed the Michael addition reaction of pyrimidine with disaccharide acrylate in
pyridine at 50 ºC for 72 hours to obtain the final products in yields from 56 to 75%. In
addition, the study of hydrolase-catalyzed Michael addition of imidazole derivatives to
acrylic monomers in organic medium has also been investigated [323]. A variety of
hydrolases were used as catalysts in this study and the reactions were conducted in
organic solvents at 50 ºC for 24 hours. All hydrolases were found to be able to catalyze
this Michael addition reaction and lipase M showed to be the most efficient hydrolase
with the percent conversion close to 100% after 24 hours. Figure 86 illustrates some
results of this study.
Figure 86. Michael addition of imidazole and methyl acrylate catalyzed by a variety of hydrolases.[323]
N
HN
O
O
+Hydrolase
tetrahydrofuran50 oC, 24 h.
O
O
N
N
Hydrolase Conversion (%)
None Not detectedAlkaline protease from Bacillus subtilis 93.0Proteinase from Aspergillus oryzae 81.9Lipase B acrylic resin from Candida Antarctica 88.7Lipase from Candida cylindracea 86.0Amano Lipase M, from Mucorjavanicus 96.0
123
Berglund et al. exhibited the possibility of ultilizing the mutant, C. Antarctica B
Ser105Ala, to catalyze the Michael addition of thiol and amine nucleophiles to α,β-
unsaturated carbonyl compounds in organic solvent [324]. The mutant enzyme was
designed by the substitution of Ser 105 to Alanine in the active-site of C. Antarctica
lipase B. This mutation led to a change in the catalytic mechanism of the enzyme.
According to turnover numbers from kinetic studies, the Ser105Ala mutant of C.
Antarctica lipase B was more efficient than the wild-type enzyme, C. Antarctica lipase B,
for the catalysis of the Michael type reaction. Recently, they also studied the use of this
Ser105Ala mutant of C. Antarctica lipase B in the catalysis of carbon-carbon bond
formation between 1,3-dicarbonyls and α,β-unsaturated carbonyl compounds (Figure 87)
[325]. The ability of wild-type and Ser105Ala mutant of C. Antarctica lipase B to
catalyze this Michael reaction was investigated under solvent free conditions. The results
showed that the reactions proceeded approximately 1.3 to 830 times faster with the
mutant than with the wild-type enzyme. In addition, the uncatalyzed reaction, without
enzyme, demonstrated a very low reaction rate. This indicates that the enzyme catalyzed
the Michael addition reactions.
Figure 87. Michael addition of acetylacetone to acrolein catalyzed by a C. Antarctica lipase B Mutant.[325]
O O
+H
O
H
O O
O
Candida antarctica lipase BSer105Ala
20 oC
100% conversion (in less than 10 minute)
124
CHAPTER 3
EXPERIMENTAL MATERIALS AND PROCEDURES
3.1 Materials
3.1.1 Chemicals
All chemicals, except 2-methoxyhydroquinone, were obtained from Aldrich. 2-
Methoxyhydroquinone was obtained from TCI America. All chemicals were used as
received without further purification. Solvents, including ethyl acetate, hexane, petroleum
ether, and acetone, were obtained from VWR and used as received without further
purification. Water in all experiments was deionized water.
3.1.2 Enzymes
Laccase (EC 1.10.3.2) used in this study was donated by Novozymes
(Franklinton, North Carolina). The laccase (NOVO NS51002) was isolated from the
white-rot fungus Trametes villosa and expressed in an Aspergillus host. Lipases were
purchased from Aldrich. Unit definition of each lipase is different depending on the
method that Aldrich used to measure lipase activity. All enzymes were kept frozen until
use.
3.1.2.1 Enzyme Assay
Laccase activity was determined by oxidation of 2,2’-azinobis-(3-ethylbenzyl thiozoline-
6-sulphonate) (ABTS) [326].The assay mixture contained 25 μM ABTS, 0.10 M sodium
125
acetate (pH 5.0), and a suitable amount of enzyme. The oxidation of ABTS was followed
by an absorbance increase at 420 nm (ε420 = 3.6 x 104 M-1cm-1) (see Figure 88 and
Figure 89). Enzyme activity was expressed in units (U = μmol of ABTS oxidized per
minutes).
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5
Time (min)
Absorba
nce
Figure 88. Graph illustrates the absorbance increase of laccase-oxidized ABTS at 420 nm.
Figure 89. Picture illustrates the changing in color of ABTS (in water) after adding laccase. The color changes from bright green to dark green.
126
3.1.3 Pulp
A commercial linerboard softwood kraft pulp (17% of lignin content, kappa
number is 113) was obtained from a southeastern U.S.A manufacturing facility. The
lignin content of the kraft pulps was determined by KMnO4 titration of the pulp following
TAPPI method T-236 [327] and expressed as a “kappa number”. This value is an indirect
measurement of lignin content (% lignin content = 0.15 x kappa number). The pulp was
exhaustively washed until the filtrate was pH neutral and colorless. Pulp was air dried
and soxhlet extracted (see Figure 90) for 24 hours with acetone with subsequent washing
with water prior to all treatments.
Figure 90. Photograph of the equipment set for soxhlet extraction.
127
3.2 Experimental Procedures for the Use of Laccase
in Organic Synthesis
3.2.1 General Information
All reactions were monitored by TLC. TLC was performed on aluminum sheets
precoated with silica gel 60 F254 (EMD Chemicals). Melting point was measured using
electrothermal MEL-TEMP instrument.
Column chromatography was performed on Combiflash Companion instrument
(Teledyne Isco company) (Figure 91). The Combiflash Companion is a flash
chromatography system which provides a fully automated system from solvent injection
to product collection. Columns used with this instrument are pre-packed columns
(RediSep columns). RediSep normal-phase silica flash columns were used in this study.
The column size is 12 g or 40 g, depending on sample size. The linear gradient elution
was used to separate mixture of the products and the flow rate is 25 – 40 ml/min.
Figure 91. Picture of Combiflash Companion instrument (Teledyne Isco company) with 40 g RediSep normal-phase silica flash columns
128
3.2.2 Analytical Analysis Procedures
3.2.2.1 1H-NMR Characterization of the Products
1H-NMR spectra were recorded on a Bruker Advance/DMX-400 instrument
operating at 400 MHz. The qualitative 1H experiments were performed using using a 90o
pulse and 3.0 s delay. The acquisitions were performed at room temperature with 24 –
120 scans and a 1 Hz line broadening.
3.2.2.2 13C-NMR Characterization of the Products
13C NMR spectra were recorded on a Bruker Advance/DMX-400 instrument
operating at 100 MHz. Acquisition was performed using a 90o pulse with a gate-
decoupling pulse sequence and 2.0 s delay between repetitions. The acquisitions were
performed at room temperature with 400 - 4000 scans and a 10 Hz line broadening.
3.2.2.3 Fourier Transform Infrared (FTIR) Spectroscopy
Fourier Transform Infrared (FTIR) transmission spectra were collected for each of
the samples in the solid state using a Magna-IR System 550 (Nicolet Instrument
Corporation). Number of scans was 64 for each sample. Pellets were formed by pressing
mixtures of 3 mg of dry sample and 500 mg of dry spectroscopy grade potassium
bromide (KBr) at 15000 psi for 3 min. under vacuum.
3.2.2.4 Mass Spectroscopy
Mass and high resolution mass spectra were carried out in The Georgia Institute
of Technology Bioanalytical Mass Spectrometry Facility. The mass analysis was
129
performed in VG instruments 70SE. The VG-70SE is capable of high resolution (~
50,000 at 10% valley) and accurate mass measurement (< 5 ppm) analyses. It is equipped
with a dedicated GC and is capable of ionization via electron impact (EI) and chemical
ionization (CI) for analysis of low molecular mass (< 700 Da), non-polar, volatile
molecules.
3.2.2.5 UV/Vis for Enzyme Activity Measurement
Laccase activity was measured using a Perkin-Elmer Lambda 900 UV/vis
spectrometer (Perkin Elmer, Waltham, MA, USA) equipped for measuring liquid
samples. The Ultraviolet-visible (UV/vis) absorbtion spectra were scan at 420 nm for 5
minutes. The example spectrum is shown in Figure 88.
3.2.3 General Procedure of the Synthesis of 1,4-Naphthoquinones and Related
Structures. (Chapter 4)
Oxygen was bubbled to a stirred solution of 30 ml of 0.10M acetate buffer (pH
4.5) and laccase (100 U) at 70 °C for 30 minutes. Next, p-hydroquinone (1.00 mmol) and
diene (2.00 mmol) were added into the reaction mixture, and stirred under air, at 70 °C
(Figure 92). In the first three hours of the reaction, 100 U of laccase was added each hour.
After 24 hours of the reaction, the reaction mixture was extracted by EtOAc (3 x 30 ml).
The organic phase was combined, dried over MgSO4, and evaporated. The resulting
crude products were purified by Combiflash Companion instrument using Redisep
normal-phase silica column. Ethyl acetate and hexane (linear gradient: 0 – 30% EtOAc)
130
were used as an eluent to obtain the products. Products were characterized by 1H-NMR,
13C NMR and MS.
Figure 92. The reaction setting of the synthesis of 1,4-naphthoquinones and related structures via laccase-catalyzed Diels-Alder reaction. 3.2.4 General Procedure of the Synthesis of o-Naphthoquinones. (Chapter 5)
In a 250-mL round-bottom flask, 20 ml of cold 0.10M acetate buffer pH 4.5 and
diene (10.00 mmol) were mixed together. The flask was then placed in an ice bath over
a stirring plate. Next, 1.00 mmol of catechol dissolved in 20 mL of 0.10M acetate buffer,
and laccase (100U) were added to the flask drop-wise. In the next three hours of the
reaction, 100 U of laccase was added each per hour.The reaction was then stirred under
room temperature. After 24 hours of the reaction, the reaction mixture was extracted by
EtOAc (3 × 30 ml). The organic phase was combined, dried over MgSO4, and
evaporated. The resulting crude products were purified by Combiflash Companion
131
instrument using Redisep normal-phase silica column. Ethyl acetate and petroleum ether
(linear gradient: 0 – 30% ethyl acetate) were used as an eluent to obtain the product.
Products were characterized by 1H-NMR, 13C NMR and MS.
3.2.5 General Procedure of the Synthesis of Benzofuran Derivatives via Laccase-
Oxidation-Michael Addition. (Chapter 6)
In a 250-mL round-bottom flask, 30 ml of 0.10 M phosphate buffer pH 7.0 and
catechol (1.00 mmol) were mixed together. Next, 100 U of lacase was added to reaction
mixture and then, 1,3-dicarbonyl compound (2.00 mmol), Sc(OTf)3 (0.20 mmol, 0.0984
g), SDS (0.20 mmol, 0.0576g), and laccase (100 U) were added. The reaction was then
stirred under air at room temperature for 1-4 hours. After the reaction was finished, the
reaction mixture was then filtrated and washed with water to collect the precipitate
product. If the product did not precipitate, the reaction mixture was extracted by EtOAc
(3 × 30 ml). The organic phase was combined, dried over MgSO4, and evaporated. The
resulting crude products were purified by Combiflash Companion instrument using
Redisep normal-phase silica column. Ethyl acetate and petroleum ether (linear gradient: 0
– 20% ethyl acetate) were used as an eluent to obtain the benzofuran product. Products
were characterized by 1H-NMR, 13C NMR and MS.
132
3.2.6 General Procedure of the Synthesis of Benzofuran Derivatives Using Laccase-
Lipase Co-Catalytic System. (Chapter 7)
In a 250-mL round-bottom flask, 30 ml of 0.10 M phosphate buffer pH 7.0 and
catechol (1.00 mmol) were mixed together. Next, 100 U of laccase was added to reaction
mixture and then, 1,3-dicarbonyl compound (2.00 mmol) and 924 U of lipase PS were
added. The reaction was then stirred under air at room temperature for 4 hours. After the
reaction was completed, the reaction mixture was extracted by EtOAc (3 × 30 ml). The
organic phase was combined, dried over MgSO4, and evaporated. The resulting crude
products were purified by Combiflash Companion instrument using Redisep normal-
phase silica column. EtOAc and petroleum ether (linear gradient: 0 – 20% ethyl acetate)
were used as an eluent to obtain the benzofuran product. Products were characterized by
1H-NMR, 13C NMR and MS.
3.2.7 General Procedure for the Reaction of Catechols and Anilines Catalyzed by
Laccase-Lipase Co-Catalytic System. (Chapter 7)
In a 250-mL round-bottom flask, 30 ml of 0.10 M phosphate buffer pH 7.0 and
catechol (1.00 mmol) were mixed together. Next, 100 U of laccase was added to reaction
mixture and then, aniline (2.00 mmol) and 924 U of lipase PS were added. The reaction
was then stirred under air at room temperature for 3.5 hours. After the reaction was
finished, the reaction mixture was filtered to collect the solid red color product. Products
were characterized by 1H-NMR, 13C NMR and MS.
133
3.3 Experimental Procedures for the Use of Laccase in Fiber Modification
3.3.1 Pulp Treatment
Laccase (80 U/1g pulp) and an amino acid (3.2 mmol/1g pulp) were added with
stirring to a 5% consistency [mass pulp/(mass pulp + water)] aqueous suspension of
linerboard pulp buffered to pH 7 with 0.10 M sodium phosphate solution. The resulting
slurry was stirred for 4 h at room temperature and then left stand 20 h. After treatment,
the pulp sample was filtered, washed with deionized water until the filtrate was colorless
and air-dried. Typically, pulp mass recovery was 95% (on oven dried weight basis).
3.3.2 Bulk Acid Group Measurement
Conductrometric titration for bulk acids was based on the work of Katz [328]. In
brief, pulp (1.50 g o.d.) was stirred in 300.00 ml of 0.10 M HCl for 1 hour followed by
rinsing in a fine fritted funnel with deionized water. The sample was then re-suspended in
250.00 ml of 1 mM NaCl solution, spiked with 1.50 ml of 0.10 M HCl and titrated
against 0.05 M NaOH at 0.25 ml increments in an atmosphere of nitrogen, recording the
conductivity at each increment. The titration data was plotted as conductivity vs. volume
of NaOH to determine the milli-equivalent of acid groups per g of pulp (Figure 93).
Trend lines were added in Excel in order to draw lines through each linear region on the
graph. A line across the “flat” portion of the curve was plotted too. The intersections of
the left trendline and the right trendline with the flat line were obtained, and their X-axis
values are represented by A and B (Figure 93). The carboxylic acid content of pulp fibers
134
is obtained using Equation 3. The reported results were the average of two measurements
which typically differed by less than 3%.
RCOOH content = (B – A) × 5 mmol/ 100 g o.d. pulp w where w is the oven dried (o.d.) weight of the pulp sample in grams.
Equation 3. The equation used to calculate for the carboxylic content of pulp fibers.
Acid Group Content Measurement
y = -0.0758x + 0.5237R2 = 0.9982
y = 0.0427x - 0.1114R2 = 0.9965
0
0.1
0.2
0.3
0.4
0.5
0.6
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
Volume of NaOH (ml)
Con
duct
ivity
Figure 93. The titration data plotted as conductivity vs. volume of NaOH for the calculation of carboxyl group (RCOOH) content using conductivity method.
A B
135
3.3.3 Pulp Refining and Handsheet Formation
Treated pulps and control were disintegrated for 30,000 revolutions (Figure 94)
and then were refined in a PFI mill (Figure 95) for 3,000 revolutions according to TAPPI
Standard T 248 [327]. Handsheets were formed according to TAPPI Standard T 205
[327] (Figure 96) and TAPPI conditioned (23 ˚C, 50% relative humidity) for at least 24
hours before physical testing.
Figure 94. Picture of instrument used for pulp disintegration.
136
Figure 95. The PFI mill for the laboratory refining of pulp.
Figure 96. Handsheet making apparatus (left) and handsheet made from liner board softwood kraft pulp (right).
137
3.3.4 Paper Physical Tests
Apparent density, tensile strength, tearing resistance, and wet tensile strength
were determine according to TAPPI methods T 210, T 494, T 414, and T 456 [327].
Apparent density was measured using a Lorenzten and Wettre caliper gauge. Tensile
testing was carried out on an Lorentzen and Wettre Alwetron tensile tester, and wet
tensile testing was measured on an Instron tester connected to a data analysis system
running Test Works Software (Figure 97). Tear tests were performed on an Elmendorf
tearing tester (Figure 98).
Figure 97. Tensile testers a) an Lorentzen and Wettre Alwetron tensile tester; b) an Instron tensile tester.
138
Figure 98. An Elmendorf tearing tester. 3.3.5 Nitrogen Analysis
Nitrogen analysis was performed on oven dry samples (24 hours, 105oC) by
elemental microanalysis at Huffman Laboratories, Inc., Golden, CO. The results are
reported on a dried sample basis.
Nitrogen is determined on a Thermo Flash analyzer. The technique is the classical
Dumas method, with thermal conductivity detection. The method is described in ASTM
D5373 (coal) and ASTM D5291 (petroleum products).
Weighed samples are combusted in oxygen at 950°C. The combustion products,
including N and NOx, are swept with a helium carrier gas through combustion catalysts,
scrubbers, and through a tube filled with reduced copper. The copper removes excess
oxygen and reduces NOx to N2. The N2 is then separated from other gases on a
chromatography column and measured with the thermal conductivity detection.
139
Precision is usually given as ± 0.3% absolute or 3% relative whichever is larger.
The detection limit can be lowered by using larger samples. For organic materials
0.02% can be obtained. Lower detection levels can be obtained for samples consisting
largely of inert materials such as soils.
3.3.6 Scanning Electron Microscope (SEM) The SEM pictures of handsheets were taken using a Hitachi S-800 FE-SEM. The
handsheet sample was stuck on the SEM sample holding stub by the conductive double
side sticky carbon film and then was coated with alloy of Au/Pt prior to analysis.
140
CHAPTER 4
ONE-POT SYNTHESIS OF 1,4-NAPHTHOQUINONES AND
RELATED STRUCTURES WITH LACCASEi
4.1 Introduction
The most abundant and available resource on the planet, one in which
biochemical processes take place, is the aqueous medium, water. Recently, water has
begun to be regarded as an environmentally friendly solvent in organic chemistry. In
addition to its environmental benefits, the use of water as a solvent is both inexpensive
and safe. In recent decades, the study of the organic reactions in aqueous solvent has
accelerated and many, often, surprising discoveries have been made [36-38,41,329].
Breslow and Rideout [35] were the first to show the beneficial effects of water on the
reactivity and selectivity of Diels-Alder reaction, quantitatively. This discovery
stimulated further research in this area. Shortly after, several studies showed that many
chemical reactions (such as pericyclic, condensation, oxidation, and reduction reactions)
could be conducted efficiently in the aqueous medium [41,330-335].
Among the organic reactions investigated in the aqueous medium, the most
widely studied reaction is the Diels-Alder reaction [34,43], a powerful tool frequently
i This manuscript was published in [Green Chemistry, 2007, 9, 475-480]- Reproduced by permission of The Royal Society of Chemistry (RSC). It is entitled as “One-pot synthesis of 1,4-naphthoquinones and related structures with laccase”. The other author is Dr. Arthur J. Ragauskas from the School of Chemistry and Biochemistry at the Georgia Institute of Technology.
141
employed to synthesize six-membered ring systems and one of the most useful reactions
for introducing structural complexity in (total) synthesis [336-339]. The Diels-Alder
reaction has many useful variations, one of which is its use in the synthesis of
anthraquinones and naphthoquinones [340,341]. Naphthoquinones have attracted
considerable attention in total synthesis because of their wide spectrum of biological
activities, such as antitumor [342,343], wound healing [344], anti-inflammatory [344],
and antimicrobial [345] and antiparasitic activities [346,347]. Another useful application
of the Diels-Alder reaction is the quinone Diels-Alder (QDA) reaction (Figure 99)
[56,61,65,348,349]. In this reaction, quinones are employed as dienophiles, which
normally possess electron-withdrawing groups. This classed of quinones are usually
unstable and difficult to isolate. To overcome these difficulties, many studies have
focused on the Diels-Alder reaction of in situ-generated quinones [350-352]. Herein, we
report the use of the enzyme, laccase, used in the in situ generation of quinones.
Laccases (benzenediol:oxygen oxidoreductase, EC 1.10.3.2) are multi-copper-
containing oxidoreductase enzymes widely distributed in plants and fungi. They are able
to catalyze the oxidation of various low-molecular weight compounds, specifically,
phenols and anilines; while concomitantly, reducing molecular oxygen to water [3-
7,149,167]. Moreover, due to their high stability, selectivity for phenolic substructures,
and mild reaction conditions used in laccase-catalyzed reactions, laccases are attractive
for fine chemical synthesis. Therefore, interest in the potential use of these enzymes in
organic synthesis has recently increased [11,13]. Indeed, a number of laccase-catalyzed
reactions has been reported [11-19]. Recently, laccase was examined in the field of
enzyme-initiated domino reaction chemistry. For example, utilizing their well known
142
propensity to oxidize phenolics, Lalk et al. [11] reported a laccase catalyzed a nuclear
animation tandem reaction. These studies have demonstrated the synthetic research
capabilities of this oxidative enzyme.
Figure 99. The Quinone Diels-Alder (QDA) reaction.
This study presents work on the synthesis of 1,4-naphthoquinones and related
structures in the aqueous medium. In this procedure, para-quinone, generated in situ from
the oxidation of para-hydroquinone by laccase, underwent the quinone Diels-Alder
reaction with a diene, and then the Diels-Alder adduct was converted directly into
dihydro 1,4-naphthoquinone. Upon extended treatment, this initial product was further
oxidized to naphthoquinone as summarized in Figure 100. The effects of a laccase dose
and temperature on these reactions, with the reaction of 2-methoxyhydroquinone (1a) and
2,3-dimethyl-1,3-butadiene (2a) as a model system, are reported here. This study also
investigated the sensitivity of this reaction system to a variety of para-hydroquinones and
dienes.
O
O
O
O
143
Figure 100. The proposed reaction pathway of laccase-catalyzed Diels-Alder reaction of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a).
4.2 Experimental Section
4.2.1 Materials
2-Methoxyhydroquinone was obtained from TCI America. Other hydroquinones,
dienes, and reagents were obtained from Aldrich. All chemicals were used as received.
Laccase (EC 1.10.3.2) from Trametes Villosa was donated by Novo Nordisk Biochem,
North Carolina.
OH
OH
MeO
1a
Laccase
O
O
MeO
2a
+
MeO
O
O
MeO
O
O
[O]
3a
[O]MeO
O
O
4a
Diels-Alder
144
4.2.2 Enzyme Assay
Laccase activity measurement is described in Chapter 3 (Experimental Materials
and Procedures).
4.2.3 General Procedure for the Study of the Effect of Laccase Dose and
Temperature
Oxygen was bubbled to a stirred solution of 30 ml of 0.10M acetate buffer (pH
4.5) and laccase (¼ of the total amount of laccase used in this reaction) at a desired
temperature for 30 minutes. Next, 2-methoxyhydroquinone (1a) (1.00 mmol) and 2,3-
dimethyl-1,3-butadiene (2a) (2.00 mmol) were added to the reaction mixture, and stirred
under air. In the first three hours of the reaction, ¼ of the total amount of laccase was
added each hour. After the reaction reached the desired reaction time, the reaction
mixture was extracted by EtOAc (3 x 30 ml). The organic phase was combined, dried
over MgSO4, and evaporated. Then the quantitative analyses of 3a and 4a were
determined by 1H-NMR spectroscopy of the crude mixture using 10 μl of
pentrafluorobenzaldehyde as an internal standard and using 0.5 ml of CDCl3 as a NMR
solvent. The example 1H-NMR spectrum is illustrated in Figure 101. Peak at 5.87 ppm
(C-H) is used to calculate yield of compound 3a and peak at 7.77 ppm (C-H aromatic) is
used to calculate yield of compound 4a.
145
Figure 101. 1H-MNR spectrum of crude mixture from the laccased-catalyzed reaction of of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a). Peaks of compound 3a are illustrated in blue boxes. Peaks of compound 4a are illustrated in red boxes. Peak of pentafluorobenzaldehyde is illustrated in green box.
4.2.4 General Procedure of the Synthesis of 1,4-Naphthoquinones and Related
Structures.
The detail of the reaction procedure is described in Chapter 3 (Experimental
Materials and Procedures).
146
4.2.5 Product Characterization
Products 3a, 3b, 3c, 3d, 4a, 4b, 4c, 4d, 4e, 4g, and 4h have been previously
reported and characterized. Compounds 3e, 3g, and 4f have also been previously reported
but without proper spectral characterization. Structures 3f and 3h are, to the best of our
knowledge, new compounds. The NMR spectra of 3f and 3h were shown in Appendix
A.1. All known products provided satisfactory analytical and spectroscopic data
corresponding to the reported literature values.
5,8-Dihydro-2-methoxy-6,7-dimethyl-1,4-naphthoquinone (3a)
Yellow crystal; mp 134-136 °C (from EtOH) (lit. [353], 133-135 °C); 1H NMR (400
MHz; CDCl3): δ 1.73 (s, 6H), 3.02 (s, 4H), 3.81 (s, 3H), 5.87 (s, 1H); 13C NMR (100
MHz, CDCl3): δ 18.1, 18.1, 30.3, 30.7, 56.1, 106.9, 121.7, 137.5, 140.2, 158.4, 181.7,
187.1; m/z (EI) 218 (M+, 33%), 216 (100), 201 (38), 187 (40), 175 (35), 159 (9), 145
(10), 117 (32), 91 (12), 69 (15), 51 (6), 39 (4); m/z (EI) 218.09211 (C13H14O3 requires
218.09429).
5,8-Dihydro-2-methoxy-5,7-dimethyl-1,4-naphthoquinone (3b)
Orange-yellow crystalline solid; mp 118-119 °C (from EtOH) (lit. [354], 118.5-120.5
°C); 1H NMR (400 MHz; CDCl3): δ 1.16 (d, J = 6.9 Hz), 1.80 (s, 3H), 2.93 (md, J =
23.4 Hz, 1H), 3.15 (md, J = 23.4 Hz, 1H), 3.29 (m, 1H), 3.81 (s, 3H), 5.47 (m, 1H),
5.88 (s, 1H); 13C NMR (100 MHz, CDCl3): δ 19.7, 20.9, 24.7, 32.8, 56.1, 106.8,
116.7, 136.3, 140.2, 142.5, 158.5, 181.4, 187.0; m/z (EI) 218 (M+, 63%), 203 (100),
175 (73), 133 (12), 119 (54), 91 (24), 69 (20), 51 (5); m/z (EI) 218.09108 (C13H14O3
requires 218.09429).
147
5,8-Dihydro-2-methoxy-5,8-dimethyl-1,4-naphthoquinone (3c)
Orange plates; mp 109-110 °C (from EtOH) (lit. [355], 110-112 °C); 1H NMR (400
MHz; CDCl3): δ 1.16 (s, 3H), 1.18 (s, 3H), 3.36-3.37 (m, 2H), 3.81 (s, 3H), 5.75 (m,
2H), 5.86 (s, 1H); 13C NMR (100 MHz, CDCl3): δ 22.5, 23.0, 29.2, 29.4, 56.0, 107.1,
128.7, 142.1, 144.6, 158.2, 181.5, 186.6; m/z (EI) 218 (M+, 52%), 203 (100), 175
(84), 133 (14), 119 (66), 91 (33), 69 (26), 39 (6); m/z (EI) 218.09397 (C13H14O3
requires 218.09429).
5,8-Dihydro-2,6,7-trimethyl-1,4-naphthoquinone (3d)
Yellow needles; mp 88-89 °C (from EtOH) (lit. [356], 87-89 °C); 1H NMR (400
MHz; CDCl3): δ 1.72 (s, 6H), 2.04(s, 3H), 2.99 (s, 4H), 6.54 (s, 1H); 13C NMR (100
MHz, CDCl3): δ 15.8, 18.2, 20.2, 30.5, 30.7, 121.8, 121.9, 133.0, 133.0, 139.5, 145.6,
187.3, 187.5; m/z (EI) 202 (M+, 100%), 187 (36), 159 (67), 119 (27), 91 (18), 39 (9);
m/z (EI) 202.10014 (C13H14O2 requires 202.09938).
5,8-Dihydro-2,5,8-trimethyl-1,4-naphthoquinone (3e)
Yellow liquid; 1H NMR (400 MHz; CDCl3): δ 1.25 (d, J = 2.2 Hz, 3H), 1.26 (d, J =
2.2 Hz, 3H), 2.10 (s, 3H), 3.43 (m, 2H), 5.83 (d, J = 2.7, 2H), 6.61(s, 1H); 13C NMR
(100 MHz, CDCl3): δ 15.7, 22.8, 22.8, 29.3, 29.5, 128.8, 128.9, 133.2, 144.0, 144.1,
145.4, 186.9, 187.3; m/z (EI) 202 (M+, 100%), 187 (79), 159 (41), 119 (56), 91 (26),
39 (7); m/z (EI) 202.09985 (C13H14O2 requires 202.09938).
1,4-Dihydro-6-methoxy-1,4-ethanonaphthalene-5,8-dione (3f)
Yellow needles; mp 123-124 °C (from EtOH); 1H NMR (400 MHz; CDCl3): δ 1.35
(d, J = 6.8 Hz, 2H), 1.49 (d, J = 8 Hz, 2H), 3.80 (s, 3H), 4.34 (br s, 1H), 4.37 (br s,
148
1H), 5.76 (s, 1H), 6.39 ( br s, 2H); 13C NMR (100 MHz, CDCl3): δ 24.4, 33.4, 33.7,
56.2, 105.9, 133.5, 133.7, 146.0, 149.2, 158.5, 178.1, 183.7; νmax/cm-1 3055, 2938,
2869, 1668, 1642, 1624, 1598, 1582, 1452, 1380, 1224, 1135, 1013, 868, 818; m/z
216 (M+, 21%), 188 (100, M - CH2CH2), 173 (39), 158 (52), 130 (14), 102 (28), 89
(33), 69 (14), 51 (8); m/z (EI) 216.08073 (C13H12O3 requires 216.07864).
1,4-Dihydro-6-methyl-1,4-ethanonaphthalene-5,8-dione (3g)
Yellow needles; mp 81-82 °C (from EtOH) (lit. [357], 83-84 °C); 1H NMR (400
MHz; CDCl3): δ 1.34 (m, 2H), 1.47 (m, 2H), 2.03 (d, J = 1.6 Hz, 3H), 4.31 (br m,
1H), 4.35 (br m, 1H), 6.38 (dd, J = 2.7 Hz, 3.8 Hz, 2H), 6.44 (q, J = 1.6 Hz, 1H); 13C
NMR (100 MHz, CDCl3): δ 15.7, 24.5, 33.5, 33.7, 132.1, 133.6, 133.7, 144.8, 148.1,
148.1, 183.9, 184.1; m/z (EI) 200 (M+, 14%), 172 (100, M - CH2CH2), 144 (27), 116
(14), 104 (18), 76 (10), 39 (3); m/z (EI) 200.08399 (C13H12O2 requires 200.08373).
1,4-Dihydro-6-bromo-1,4-ethanonaphthalene-5,8-dione (3h)
Orange crystals; mp 104-106 °C (from EtOH); 1H NMR (400 MHz; CDCl3): δ 1.39 (d,
J = 8.5 Hz, 2H), 1.53 (d, J = 8 Hz, 2H), 4.35 (br m, 1H), 4.44 (br m, 1H), 6.42 (t, J =
3.4 Hz, 2H), 7.15 (s, 1H); 13C NMR (100 MHz, CDCl3): δ 24.5, 24.6, 33.9, 34.8,
133.5, 133.7, 136.7, 137.0, 147.9, 148.9, 175.8, 181.1; ; νmax/cm-1 3043, 2998, 2935,
2869, 1660, 1645, 1627, 1571, 1445, 1331, 1302, 1263, 1233, 1051, 892, 777; m/z
(EI) 266 (M + 2, 9%), 264 (M+, 9%), 238 (82), 236 (80), 185 (10), 157 (100), 129
(41), 101 (21), 76 (11), 51 (7); m/z (EI) 263.97755 (C12H9O2Br requires 263.97859).
149
2-Methoxy-6,7-dimethyl-1,4-naphthoquinone (4a)
Yellow crystal; mp 165-167 °C (from EtOH) (lit. [358], 169-171 °C); 1H NMR (400
MHz; CDCl3): δ 2.36 (s, 6H), 3.87 (s, 3H), 6.06 (s, 1H), 7.77 (s, 1H), 7.82 (s, 1H);
13C NMR (100 MHz, CDCl3): δ 19.9, 20.2, 56.3, 109.5, 127.1, 127.6, 128.9, 129.9,
142.9, 144.2, 160.2, 180.2, 185.1; m/z (EI) 216 (M+, 100%), 201 (33), 187 (38), 186
(27), 158 (8), 145 (8), 130 (15), 117 (25), 103 (5), 91 (5), 69 (4), 51 (4), 39 (3); m/z
(EI) 216.07925 (C13H12O3 requires 216.07864).
2-Methoxy-5,7-dimethyl-1,4-naphthoquinone (4b)
Yellow powder; mp 149-151 °C (from EtOH) (lit. [354], 146-148 °C); 1H NMR (400
MHz; CDCl3): δ 2.40 (s, 3H), 2.62 (s, 3H), 3.87 (s, 3H), 6.07 (s, 1H), 7.45 (s, 1H),
7.85 (s, 1H); 13C NMR (100 MHz, CDCl3): δ 17.0, 21.2, 56.2, 107.9, 124.1, 128.6,
131.6, 134.8, 140.3, 144.2, 160.8, 181.9, 184.7; m/z (EI) 216 (M+, 100%), 201 (73),
186 (8), 128 (7), 117 (27), 103 (4), 91 (4), 77 (5), 63 (4), 51 (4), 39 (2); m/z (EI)
216.07918 (C13H12O3 requires 216.07864).
2,6,7-Trimethoxy-1,4-naphthoquinone (4c)
Golden yellow solid; mp 232-234 °C (lit. [359], 234-235 °C); 1H NMR (400 MHz;
CDCl3): δ 3.90 (s, 3H), 4.04 (s, 6H), 6.07 (s, 1H), 7.51 (s, 1H), 7.54(s, 1H); 13C NMR
(100 MHz, CDCl3): δ 56.3, 56.5, 107.7, 108.1, 109.0, 125.4, 126.9, 152.9, 153.8,
160.3, 179.4, 184.5; m/z (EI) 248 (M+, 100%), 233 (10), 219 (37), 205 (6), 177 (17),
162 (10), 149 (28), 134 (6), 119 (6), 93 (3), 69 (6), 63 (3); m/z (EI) 248.06705
(C13H12O5 requires 248.06847).
150
2-Methoxy-1,4-naphthoquinone (4d)
Yellow solid; mp 179-182 °C (lit. [360], 178-182 °C); 1H NMR (400 MHz; CDCl3): δ
3.93 (s, 3H), 6.18 (s, 1H), 7.71-7.75 (m,2H), 8.06-8.13 (m, 2H); 13C NMR (100 MHz,
CDCl3): δ 56.4, 109.8, 126.1, 126.6, 130.9, 131.9, 133.3, 134.3, 160.4, 180.0, 184.7;
m/z (EI) 188 (M+, 100%), 173 (40), 158 (36), 102 (40), 89 (52), 76 (20), 69 (10), 50
(10), 39 (2); m/z (EI) 188.04625 (C11H8O3 requires 188.04734).
2,6,7-Trimethyl-1,4-naphthoquinone (4e)
Yellow solid; mp 104-105 °C (lit. [361], 105-106 °C); 1H NMR (400 MHz; CDCl3): δ
2.18 (s, 3H), 2.40(s, 6H),6.77 (s, 1H), 7.78 (s, 1H), 7.82 (s, 1H); 13C NMR (100 MHz,
CDCl3): δ 16.4, 16.4, 20.1, 127.0, 127.5, 130.1, 130.2, 135.4, 143.3, 143.3, 147.8,
185.3, 185.7; m/z (EI) 200 (M+, 100%), 185 (11), 172 (36), 157 (16), 144 (9), 132 (
27), 115 (4), 104 (10), 77 (6), 63 (4), 51 (5), 39 (4); m/z (EI) 200.08187 (C13H12O2
requires 200.08373).
2-Methyl-6,7-dimethoxy-1,4-naphthoquinone (4f)
Orange-yellow solid; mp 211-212 °C (lit. [362], 211-212.5 °C); 1H NMR (400 MHz;
CDCl3): δ 2.16 (s, 3H), 4.01 (s, 6H), 6.73 (s, 1H), 7.47 (s, 1H), 7.51 (s, 1H); 13C NMR
(100 MHz, CDCl3): δ 16.5, 56.5, 56.5, 107.5, 108.0, 111.4, 126.9, 127.1, 135.2,
147.7, 153.2, 184.6, 185.1; m/z (EI) 232 (M+, 100%), 202 (31), 189 (19), 136 (12), 93
(7), 39 (8); m/z (EI) 232.08528 (C13H12O4 requires 232.07356).
2-methyl-1,4-naphthoquinone (menadione) (4g)
Bright yellow solid; mp 104-105 °C (lit. [360], 103-104 °C); 1H NMR (400 MHz;
CDCl3): δ 2.17 (s, 3H), 6.83 (s, 1H), 7.71-7.73 (m, 2H), 8.03-8.09 (m, 2H); 13C NMR
151
(100 MHz, CDCl3): δ 16.4, 126.0, 126.4, 132.0, 132.1, 133.5, 133.5, 135.5, 148.1,
184.9, 185.4; m/z (EI) 172 (M+, 100%), 144 (23), 115 (24), 104 (34), 76 (22), 50 (9);
m/z (EI) 172.05149 (C11H8O2 requires 172.05243).
2-Bromo-6,7-dimethyl-1,4-naphthoquinone (4h)
Yellow solid; mp 156-158 °C (lit. [363], 156-159 °C); 1H NMR (400 MHz; CDCl3): δ
2.42 (s, 6H), 7.45 (s, 1H), 7.82 (s, 1H), 7.91(s,1H); 13C NMR (100 MHz, CDCl3): δ 20.2,
127.8, 128.8, 129.6, 139.9, 140.1, 144.1, 144.5, 177.9, 182.6; m/z (EI) 266 (M+ + 2,
77%), 264 (M+, 77%), 185 (100), 157 (53), 128 (25), 103 (7), 77 (9), 51 (9), 39 (3); m/z
(EI) 263.97489 (C12H9O2Br requires 263.97859).
152
4.3 Results and Discussion
4.3.1 Preliminary Study of the Reaction System
Laccase-catalyzed reaction of 1,4-benzoquinones and dienes was initially
investigated by using 1a and 2a as the model reagents and laccase as an oxidizing
agent. Laccase first converted 1a to 2-methoxy-1,4-benzoquinone, and then the
quinone reacted with diene 2a via the Diels-Alder reaction. The Diels-Alder adducts
then underwent further oxidation to generate 5,8-dihydro-2-methoxy-6,7-dimethyl-
1,4-naphthoquinone (3a) and 2-methoxy-6,7-dimethyl-1,4-naphthoquinone (4a).
In this preliminary study, the total amount of laccase used in the reaction was
1000 U/ 1g substrate, and the equivalence ratio of 2-methoxy hydroquinone and 2,3-
dimethyl-1,3- butadiene was 1:2, to enhance the likelyhood that no in situ-generated
2-methoxy benzoquinone remained to further oxidize the Diels-Alder adducts. The
reaction was conducted in 0.10M acetate buffer pH 4.5, in the presence of oxygen at
50 °C, for 24 hours (Figure 102). A pH of 4.5 was chosen for this reaction system
because many studies have shown that this pH is the optimum pH for laccase activity
in the formation of quinone, as in the work of Ishihara, and Leonowicz et al. and
Ragauskas [364-366]. In this reaction system, vigorous stir was required to disperse
2a, which is slightly dissolved in water, in an emulsion to increase the reaction rate
between the in situ-generated quinone and 2a. Moreover, the hydrophobic interactions
between relatively apolar quinone and 2a forced them into close proximity and favour
the Diels-Alder reaction products.
153
Figure 102. The preliminary reaction system for laccase-catalyzed aqueous Diels-Alder reaction of 2-methoxyhydroquinone (1a) and 2,3-dimethyl-1,3-butadiene (2a).
In the preliminary study, we examined the effect of oxygen on the formation
of the products. We found that the quantity of oxygen affected the reaction. When an
excessive amount of oxygen, such as bubbling oxygen throughout the reaction or
pressurizing with oxygen at 9.9974 × 105 N/m2 (145 psi), was used, the main product
was 2-methoxy-1,4-benzoquinone (26%) and very small amounts of 3a and 4a were
generated. In contrast, stirring the reaction under air generated 3a (13%) and 4a
(45%). However, we also found that bubbling oxygen for 30 minutes into a
laccase/buffer solution before adding all the reagents and gradually adding ¼ of the
laccase (250 U/1g substrate) at the beginning of each of the first four hours of the 24-
hour reaction improved the yield of 3a and 4a to 15% and 50%, respectively. After
OH
OH
MeO
O
MeO
O
O
O
MeO1a 2a
3a
4a
Laccase, O2
0.1M Acetate Buffer pH 4.550 oC, 24 hrs
154
this reaction procedure was examined, the control reaction adding no laccase was
studied. The result showed that when no laccase was added to the system, no desired
products were obtained. Therefore, the oxidizing agent, laccase, must be added to
generate 2-methoxy-1,4-benzoquinone in situ. This quinone then underwent further
reaction with diene to generate 3a and 4a.
4.3.2 The Effect of Laccase Dose
After the preliminary study, the next approach was to optimize the reaction
conditions. The optimization was studied by investigating the effects of laccase dose
and temperature. The laccase doses used in these experiments were 500, 1000, 2000,
and 4000 U/ 1 g of 1a. The reaction was conducted at 50 °C. The method for this
study is described in the experimental section. The quantitative study of 3a and 4a
was measured by 1H-NMR spectroscopy using tetrafluorobenzaldehyde as an internal
standard. Figure 103 shows the results of the study.
155
Figure 103. The effect of laccase dose on the formation of compound 3a and 4a. The percent yield of 3a and 4a was measured by 1H-NMR spectroscopy.
156
According to Figure 103, the amount of laccase in the reaction affected the
formation of 3a and 4a. When the amount of laccase used in the reaction was increased
from 500 to 4000 U/ 1g substrate, the percent yield of 3a and 4a at the end of the reaction
(24 hours) also increased from 15% to 34% and from 47% to 60%, respectively. In
addition, the formation of 3a sharply increased in the first three hours, and then decreased
gradually throughout the reaction. In contrast, the formation of 4a increased slightly in
the first two hours and then gradually increased after the third hour of the reaction. The
explanation of this result is that the formation of 3a was predominant at the beginning of
the reaction, and upon further treatment, some of 3a was gradually oxidized to generate
4a, leading to the continual increase in the yield of 4a at the longer reaction time. The
proposed reaction pathway is summarized in Figure 100. The first step was the oxidation
of 1a by laccase to form 2-methoxy benzoquinone and then this quinone underwent the
Diels-Alder reaction with 2a to generate the Diels-Alder adduct. The Diels-Alder adduct
was then oxidized by either laccase or quinone in the reaction solution to generate 3a and
upon further treatment, 3a was oxidized to 4a. To confirm the proposed reaction
pathway, we stirred 3a in 0.10M acetate buffer, pH 4.5 at 70 ˚C for 24 hours, with either
laccase (4000 U/1g of 3a) or with 2-methoxybenzoquinone (model quinone) (1equiv.).
The results show that the percent conversion of 3a to 4a was 35% and 16% with laccase
and 2-methoxybenzoquinone respectively. Therefore, these results show that both laccase
and quinone in the reaction solution can oxidize 3a to generate 4a. However, laccase
appears to be a better oxidizing agent than the quinone in this reaction system.
157
4.3.3 The Effect of Temperature
As demonstrated in the previous section, laccase dose has an influence on the
formation of compounds 3a and 4a. The more laccase we used, the more products we
obtained. Another factor that should affect the reaction is temperature. Thus, the
experiment was conducted at different temperatures, including 25 °C, 50 °C, 70 °C,
and 100 °C. The reaction procedure was the same as that used before except 4000 U/
1g of 1a was used. Figure 104 illustrates the effect of temperature on the reaction.
It is obvious that when 4a was formed, its yield increased when the
temperature of the reaction increased. For example, at the end of the reaction, the
percent yield of 4a was 17%, 60%, and 87% for the reaction at 25 °C, 50 °C, and 70
°C, respectively. In contrast, the formation of 3a exhibited a different response to
temperatures. For the reaction at 50 °C and 70 °C, the amount of 3a sharply increased
in the first two hours, and then decreased after the second hour. However, the
decrease at 70 °C was faster than that at 50 °C because a higher temperature can more
easily accelerate the conversion of 3a to 4a. For the reaction at 25 °C, the formation
of 3a gradually increased throughout the reaction. Moreover, we found that 2-
methoxy-6,7-dimethyl-4a,5,8,8a-tetrahydro-1,4-naphthoquinone, the Diels-Alder
adduct, was the main product of the reaction at 25 °C, instead of 3a and 4a.
Therefore, this reaction best underwent the quinone Diels-Alder reaction to generate
the Diels-Alder adduct at a low temperature, and upon further treatment, the Diels-
Alder adduct was slowly converted to 3a, and only a small amount of 4a was
obtained. For the reaction at 100 °C, no products were obtained because, at this high
temperature, laccase was denatured.
158
Figure 104. The effect of temperature on the formation of compound 3a and 4a. The percent yield of 3a and 4a was measured by 1H-NMR spectroscopy. (No products were obtained at 100 °C.)
159
4.3.4 The Reaction of p-Hydroquinones and Dienes
From the optimization experiments, we chose to conduct the reaction with
4000 U of laccase/1g substrate at 70 °C for 24 hours to investigate the reaction of
various para-hydroquinones and dienes as shown in Table 9.
In this experiment, three different p-hydroquinones, in which R1 represented
the OMe, Me, and Br groups, were used and conducted with a variety of dienes. The
data in Table 9 shows that in most cases, 1,4-naphthoquinone products (4) were
obtained as major products, and only small amounts or none of dihydro 1,4-
naphthoquinone products (3) were obtained. However, when dienes have alkyl groups
at R2 and R5 (e.g., 2c and 2f), only 3-type products were formed. In addition, when R1
is the OMe group, the yield of products was higher than when R1 is the Me or Br
groups. Although quinones with a Br substituent, a strong electron-withdrawing
group, have been proven to be very reactive dienophiles for the Diels-Alder reaction,
it produced a lower yield of the products than quinones with an OMe substituent, an
electron-donating group. This result can be explained by the substrate affinity of
laccase, which varies, depending on the substituents and their recipocal positions on
the aromatic ring. Therefore, p-hydroquinones that have higher affinity to laccase are
more easily oxidized, and generate higher amounts of the starting quinone that react
with diene in the first step of the reaction. In this case, p-hydroquinones with the Br
substituent have lower affinity to laccase than p-hydroquinones with OMe. This result
agrees with that of a study that reported the high affinity of the phenolic compounds
bearing the methoxyl group to laccase [367]. In addition, substituents also have effect
on redox potential of hydroquinone starting material. Xu [198] showed that the
160
electron-withdrawing substituents reduce the electron density at the phenoxy group
and increase redox potential of molecule, thus making it more difficult to be oxidized
and less reactive in surrendering electron to the substrate pocket in laccase. In
contrast, the presence of the electron-donating substituents results in the reduction in
redox potential. Therefore, in this study, p-hydroquinone with the OMe group is more
easily oxidized than that with the Br group.
161
Table 9. The reaction of p-hydroquinones and dienesa.
p-Hydroquinone Diene Yield of Products (%)b
1a: R1 = OMe 2a: R2 = R5 = H, R3 = R4 = Me 3a (10%) 4a (60%)
1a 2b: R2 = R4 = H, R3 = R5 = Me 3b (9%) 4b (55%)
1a 2c: R3 = R4 = H, R2 = R5 = Me 3c (46%) -
1a 2d: R2 = R5 = H, R3 = R4 = OMe - 4c (12%)
1a 2e: R3 = R4 = R5 = H, R2 = OMe - 4d (79%),
R2 =H 1b: R1 = Me 2a 3d (20%) 4e (22%)
1b 2c 3e (19%) -
1b 2d - 4f (4.28%)c
1b 2e - 4g (40%), R2 = H
1c: R1 = Br 2a - 4h (21%)
1a 2f:
3f (64%)
-
1b 2f 3g (49%) -
1c 2f 3h (51%) -
aReaction conditions: The reaction of p-hydroquinone (1 equiv.) and diene (2 equiv.) was stirred with laccase (4000U/1g substrate) in 0.10M acetate buffer pH 4.5 at 70 ˚C for 24 hours. bYield of isolated products was calculated base on the amount of 1,4-benzoquinone starting materials. cFound 26% of methylbenzoquinone as another product.
OH
OH
R1
1
R2R3
R4R5
2
O
O
R1
R2R3
R4R5
3
R1
R2R3
R4R5
O
O4
162
4.4 Conclusions
Here, we reported a new green chemistry synthesis of 1,4-naphthoquinones
and related structures by using both a nonhazardous oxidizing agent, laccase, and the
environmentally benign solvent, water. This study also shows another potential use of
laccase as an oxidizing agent in organic synthesis. Moreover, the reaction system we
used in this study produced the 1,4-naphthoquinone products in high yield. However,
the reactivity of the reaction depends on the substrate specificity of laccase and the
reactivity of both generated quinones and dienes. For instance, the presence of the
electron-donating substituents, such as OMe group, results in the reduction in redox
potential and makes p-hydroquinone more easily oxidized. Therefore, in this study,
methoxy-hydroquinone provided higher yield of the product than methyl- or bromo-
hydroquinone. In addition, both temperature and laccase dose effect on the formation
the corresponding products. Therefore, the reaction condition have to be controlled to
obtain the desired products.
163
CHAPTER 5
LACCASE-GENERATED QUINONES IN 1,2-NAPHTHOQUINONE
SYNTHESIS VIA DIELS-ALDER REACTIONii
5.1 Introduction
The combination of enzymatic with nonenzymatic transformations for tandem
reactions was first reported by Waldmann and co-workers in 1998 [117]. They reported
the synthesis of highly functionalized bicycle[2.2.2]octenes by a tyrosinase-initiated
hydroxylation-oxidation of phenols followed by a Diels-Alder (DA) reaction with
electron rich dienophiles (see Figure 105). These studies, conducted in chloroform,
provided a unique three-step one-pot reaction of bicyclic DA products in high yields with
the key intermediate being reactive ortho-quinones. The applicability of enzyme
catalyzed domino reactions in green chemistry has only recently been fully appreciated.
ii This manuscript was published in Tetrahedron Letters, 2007, 48, 2983-2987. It is entitled as “Laccase-generated quinones in naphthoquinone synthesis via Diels-Alder reaction”. The other authors are Abdullah Zettili from Department of Physical and Earth Science, School of Chemistry at Jacksonville State University and Dr. Arthur J. Ragauskas from the School of Chemistry and Biochemistry at the Georgia Institute of Technology. This chapter is reproduced with the kind permission of from [Tetrahedron Letters]. Copyright © 2007 Elsevier Science.
164
Figure 105. The example of enzyme-initiated reaction cascade reported by Waldmann and co-workers.[117]
In the previous Chapter, the successful synthesis of p-naphthoquinones via
laccase-catalyzed Diels-Alder reaction between in situ-generated p-quinones and dienes
in aqueous medium was described. To broaden substrate spectrum for this laccase-
catalyed Diels-Alder reaction system, this Chapter further investigated the use of this
reaction system for the synthesis of o-naphthoquinones. In this study, a series of
substituted o-naphthoquinones were synthesized via an aqueous cascade reaction between
acyclic dienes and in-situ generated o-quinones. The ortho-quinones were synthesized in
situ by the oxidation of the corresponding o-benzohydroquinone by laccase. The initial
Diels-Alder adduct was shown to undergo further oxidization by laccase and/or quinone
to yield the desired o-naphthoquinones (see Figure 106).
OH
R1
OH
OH
R1
O
O
R1
Tyrosinase
CHCl3, O2
Tyrosinase
CHCl3, O2
R2
O
O
R1R2
O
O
R1
R2
+
165
Figure 106. Laccase-initiated cascade synthesis of substitute o-naphthoquinones via aqueous Diels-Alder reaction.
Therefore, this Chapter summarizes our interests in the use of laccase for the
synthesis of substituted o-napthoquinones. Naphthoquinones are naturally occurring
compounds which have attracted interest in total synthesis because of their wide range of
biological activity including antitumor [342,343,368], wound healing [344], anti-
inflmmatory [344], and antimicrobial [345] and antiparasitic activities [346,347].
O H
O H
R 1
R 2
R3
R4
O
O
R2
R3
R4
R5
R 1
O
O
R 1
Diels-Alder
[O]
+Laccase
0.1 M Acetate Buf f er pH 4.5
166
5.2 Experimental Section
5.2.1 Enzyme Assay
Laccase activity measurement is described in Chapter 3 (Experimental Materials
and Procedures).
5.2.2 General Procedure of the Synthesis of o-Naphthoquinones
The detail of the reaction procedure is described in Chapter 3 (Experimental
Materials and Procedures).
5.2.3 Typical Experimental Procedure for p-Naphthoquinone Synthesis
p-Hydroquinone (1.00 mmol), 1-acetoxy-1,3-butadiene (2.00 mmol), and laccase
(100 U) were stirred in 40 ml of 0.10M acetate buffer pH 4.5 under air at 55 °C. In the
next three hours of the reaction, 100 U of laccase was added each per hour. After 24
hours of the reaction, the reaction mixture was extracted by EtOAc (3 × 30 ml). The
organic phase was combined, dried over MgSO4, and evaporated. The resulting crude
products were purified by Combiflash Companion instrument using Redisep normal-
phase silica column. Ethyl acetate and petroleum ether (linear gradient: 0 – 20% ethyl
acetate) were used as an eluent to obtain the products.
167
5.2.4 Product Characterization
Most compounds have been previously reported and characterized except the two
compounds which are 4,7,8-trimethyl-1,2-naphthoquinone (6e) and 4-methyl-6,7-
dimethoxy-1,2-naphthoquinone (6f). The NMR spectra of compound 6e and 6f are shown
in Appendix A.2. All known products provided satisfactory analytical and spectroscopic
data corresponding to the reported literature values.
6,7-Dimethyl-1,2-naphthoquinone (6a)
Orange-red needles: mp 147-148 °C (lit. [369],146-147 °C); 1H NMR (CDCl3, 400 MHz)
δ 2.32 (s, 3H, CH3), 2.34 (s, 3H, CH3), 6.33 (d, J = 10 Hz, 1H, CH), 7.09 (s, 1H, Ar),
7.35 (d, J = 10 Hz, 1H, CH), 7.84 (s, 1H, Ar); 13C NMR (CDCl3, 100 MHz) δ 19.6, 20.2,
127.0, 129.5, 131.3, 131.6, 132.7, 140.1, 145.6, 146.0, 178.9, 181.3.
4,6,7-Trimethyl-1,2-naphthoquinone (6b)
Orange needles; mp. 119 °C (decomp.) (lit. [369],120 °C (decomp.); 1H NMR (CDCl3,
400 MHz) δ 2.33 (s, 3H, CH3), 2.36 (s, 3H, CH3), 2.37 (s, 3H, CH3), 6.31 (s, 1H, CH),
7.25 (s, 1H, Ar), 7.88 (s, 1H, Ar); 13C NMR (CDCl3, 100 MHz) δ 19.5, 20.6, 20.6, 126.8,
127.9, 129.1, 131.3, 133.4, 139.9, 145.4, 154.1, 179.7, 180.9; MS (EI) m/z 200 (M+,
100%), 172 (76), 157 (35), 128 (20), 91 (4), 77 (5), 51 (5); HRMS (EI) calcd for
C13H12O2 requires 200.08373, found 200.08264.
3-Methoxy-6,7-dimethyl-1,2-naphthoquinone (6c)
Maroon needles: mp 230-232 °C (lit. [369],231-233 °C); 1H NMR (CDCl3, 400 MHz) δ
2.26 (s, 3H, CH3), 2.3 (s, 3H, CH3), 3.81 (s, 3H, OCH3), 6.35 (s, 1H, CH), 6.96 (s, 1H,
Ar), 7.75 (s, 1H, Ar); 13C NMR (CDCl3, 100 MHz) δ 19.3, 20.3, 55.6, 110.5, 115.1,
168
120.9, 130.0, 131.8, 136.4, 137.2, 151.3,176.7, 178.2; MS (EI) m/z 216 (M+, 80%), 188
(100), 173 (38), 159 (34), 145 (41), 117 (93), 91 (16), 57 (11), 51 (9); HRMS (EI) calcd
for C13H12O3 requires 216.07864, found 216.07910.
4-t-Butyl-6,7-dimethyl-1,2-naphthoquinone (6d)
Orange crystals [370]: 1H NMR (CDCl3, 400 MHz) δ 1.46 (s, 9H, t-Bu), 2.27 (s, 3H,
CH3), 2.35 (s.3H, CH3), 6.38 (s, 1H, CH), 7.67 (s, 1H, Ar), 7.87 (s, 1H, Ar); 13C NMR
(CDCl3, 100 MHz) δ 19.3, 20.9, 30.9, 36.8, 124.4, 130.0, 130.8, 132.1, 138.8,144.5,
164.6, 179.8, 182.3.
4,7,8-Trimethyl-1,2-naphthoquinone (6e)
Orange solid; mp. 118 °C (decomp.); 1H NMR (CDCl3, 400 MHz) δ 2.36 (s, 3H, CH3),
2.37 (s, 3H, CH3) 2.59 (s, 3H, CH3), 6.32 (s, 1H, CH), 7.29 (d, J = 8 Hz, 1H, Ar), 7.41 (d,
J = 8 Hz, 1H, Ar); 13C NMR (CDCl3, 100 MHz) δ 17.6, 20.9, 21.2, 124.5, 126.3, 129.8,
134.8, 135.6, 141.8, 144.1, 154.9, 181.6, 183.5; MS (EI) m/z 200 (M+, 31%), 172 (100),
157 (22), 141(11), 129 (38), 115 (12), 102 (4), 77 (7), 63 (7), 51 (8), 44 (27); HRMS (EI)
calcd for C13H12O2 requires 200.0837, found: 200.0840.
4-Methyl-6,7-dimethoxy-1,2-naphthoquinone (6f)
Red needles: mp. 124 °C (decomp.); 1H NMR (CDCl3, 400 MHz) δ 2.37 (s, 3H, CH3),
3.97 (s, 3H, OCH3), 4.03 (s, 3H, OCH3), 6.25 (s, 1H, CH), 6.92 (s, 1H, Ar), 7.61 (s, 1H,
Ar); 13C NMR (CDCl3, 100 MHz) δ 20.7, 56.3, 108.7, 112.0, 125.2, 125.9, 130.8, 150.5,
153.2, 154.6, 178.3, 181.0; MS (EI) m/z 232 (M+, 54%), 204 (100), 189 (37), 175 (4), 161
(9), 133 (9), 118 (8), 105 (12), 77 (5), 63 (7), 39 (6); HRMS (EI) calcd for C13H12O4
requires 232.07356, found 232.07343.
169
4-Methyl-1,2-naphthoquinone (6g)
Orange needles: mp. 110 °C (decomp.) (lit. [371] ,109 °C (decomp.); 1H NMR (CDCl3,
400 MHz) δ 2.40 (s, 3H, CH3), 6.38 (s, 1H, CH), 7.53 (d and t, 2H, Ar), 7.70 (t, J = 8 Hz,
1H, Ar), 8.09 (d, J = 8 Hz, 1H, Ar); 13C NMR (CDCl3, 100 MHz) δ 20.5, 78.69, 126.3,
127.3, 129.7, 130.6, 131.0, 135.4, 153.7, 179.5, 180.3; MS (EI) m/z 172 (M+, 4%), 144
(100), 129 (4), 115 (71), 101 (4), 89 (7), 63 (7), 57 (6), 39 (5); HRMS (EI) calcd for
C11H8O2 requires 172.05243, found 172.05172.
1,4-Naphthoquinone (7a)
Yellow-brownish solid: mp. 127-128 °C (lit. [372] , 128 °C); 1H NMR (CDCl3, 400
MHz) δ 6.98 (s, 2H, CH), 7.75 (m, 2H, Ar), 8.07 (m, 2H, Ar); 13C NMR (CDCl3, 100
MHz) δ 126.4, 131.8, 133.9, 138.6, 185.0.
2-Methyl-1,4-naphthoquinone(menadione) (7b)
Bright yellow solid: mp. 104-105 °C (lit. [360] ,103-104 °C); 1H NMR (CDCl3, 400
MHz) δ 2.17 (s, 3 H, CH3), 6.83 (s, 1H, CH), 7.71-7.73 ( m, 2H, Ar), 8.03-8.09 (m, 2H,
Ar); 13C NMR (CDCl3, 100 MHz) δ 16.4, 126.0, 126.4, 132.0, 132.1, 133.5(x2), 135.5,
148.1, 184.9, 185.4; MS (EI) m/z 172 (M+, 100%), 144 (23), 115 (24), 104 (34), 76 (22),
50 (9); HRMS (EI) calcd for C11H8O2 requires 172.05243, found 172.05149.
2-Methoxy-1,4-naphthoquinone (7c)
Yellow needles: mp. 179-182 °C (lit. [373], 178-182 °C); 1H NMR (CDCl3, 400 MHz)
δ 3.93 (s, 3H, OCH3), 6.18 (s, 1H, CH), 7.73 (dq, J = 1 Hz and 7 Hz, 2H, Ar), 8.06 (dd, J
= 1 Hz and 7 Hz, 1H, Ar), 8.11 (dd, J = 1 Hz and 7 Hz, 1H, Ar); 13C NMR (CDCl3, 100
MHz) δ 56.4, 109.8, 126.1, 126.6, 130.9, 131.9, 133.3, 134.3, 160.4, 180.0, 184.7; MS
170
(EI) m/z 188 (M+, 100%), 173 (40), 158 (36), 102 (40), 89 (52), 76 (20), 69 (10), 50 (10),
39 (2); HRMS (EI) calcd for C11H8O3 requires 188.04734, found 188.04625.
2-Bromo-1,4-naphthoquinone (7d)
Yellow powder: mp.130-131 °C (lit. [374], 130-132 °C); 1H NMR (CDCl3, 400 MHz)
δ 7.52 (s, 1H, CH), 7.75-7.80 (m, 2H, Ar), 8.09 (m, 1H, Ar), 8.16 (m, 1H, Ar); 13C NMR
(CDCl3, 100 MHz) δ 126.8, 127.7, 130.8, 131.6, 134.0, 134.3, 140.0, 140.3, 177.8,
182.3.
2-Chloro-1,4-naphthoquinone (7e)
Yellow solid: mp. 112-113 °C (lit. [375], 112-113 °C); 1H NMR (CDCl3, 400 MHz) δ
7.23 (s, 1H, CH), 7.75-7.82 (m, 2H, Ar), 8.09 (m, 1H, Ar), 8.17 (m, 1H, Ar); 13C NMR
(CDCl3, 100 MHz) δ 126.7,127.5, 131.3, 131.7, 134.1, 134.5, 135.9, 146.3, 177.9, 182.7.
171
5.3 Results and Discussion
Initially, we focused our attention on the reaction of laccase with 1,2-catechols
yielding the corresponding o-quinones which have an interesting reactivity profile in
cycloaddition reactions [376], and have been used in o-naphthoquinone synthesis. In a
preliminary study, the reaction of catechol (5a) and 2,3-dimethyl-1,3-butadiene (2a) in
the presence of laccase was investigated. As summarized in Table 10, optimal yields of
6,7-dimethyl-1,2-naphthoquinone (6a) was achieved when the reaction was conducted
with 1 equivalence of 1 and 10 equivalence of 2 in the presence of laccase in 0.1 M
acetate buffer pH 4.5 at 3 ˚C for the first two hour of the reaction. The reaction mixture
was then warmed to room temperature and stirred for another 22 hours.
Table 10. Preliminary study of the laccase-catalyzed reaction of catechol (5a) and 2,3-dimethyl-1,3-butadiene (2a) in aqueous medium
Entry 5a : 2a (equiv.) Temperature Yielda of 6a (%)
1 1:10 3 °C (2 h), RT 47 2 1:10 RT. 10 3 1:10 60 °C No product formed 4 1:5 3 °C (2 h), RT 8 5 1:15 3 °C (2 h), RT 32 aIsolated yield.
OH
OHLaccase
0.1M acetate buffer pH 4.524 hours
O
O
2a5a 6a
172
An excess of the diene was required to overcome the intrinsic instability of the o-
benzoquinone as it will undergo competing decomposition, dimerization, and
polymerization if insufficient diene is present for the Diels-Alder reaction [369]. In
addition, the reaction temperature and medium were shown to have an effect on the
reaction outcome. For example, if the reaction was preformed at room temperature or 60
˚C the yield of 6a was diminished to only 10 and 0 %, respectively. This result was
attributed an increase in the rate of decomposition and polymerization of the in-situ
generated o-quinone. Therefore, we retarded the rate of decomposition and
polymerization by maintaining the initial reaction temperature to 3 ˚C for the first two
hours and then allowing the reaction mixture warm to room temperature. This cascade
reaction system provided 47 % of 6a. Next, we examined whether the increase of reactant
solublility by replacing acetate buffer solvent with organic or biphasic organic/water
solvent can enhance the reaction. The results of these experiments are summarized in
Table 11 . The reaction performed in aqueous acetate buffer at pH 4.5, generally known
to be the optimum pH for laccase activity in the formation of quinone [364-366],
provided the best result (see Table 11). The lower percent yield in other solvent systems
was due to a decrease of laccase activity in organic and aqueous-organic mixed solvent
[377,378]. Moreover, the Diels-Alder reaction has shown to exhibit higher reactivity and
selectivity in aqueous medium than in organic solvent [35]. Interestingly, the use of a 1:1
acetate buffer/chloroform medium, provided the aromatized DA adduct (5,8-dihydro-6,7-
dimethyl-1,2-naphthoquinone) instead of fully oxidized product (6a).
173
Table 11. Solvent effect on the laccase-catalyzed reaction of catechol (5a) and 2,3-dimethyl-1,3-butadiene (2a)a
Entry Solvent Yieldb of 6a (%)
1 0.1 M Acetate buffer pH 4.5 47
2 Water 18
3 5% Aqueous PEG 2000 25
4 p-Dioxane 0
5 1:1 p-Dioxane/acetate buffer 8
6 1:1 Ethylene Glycol/acetate buffer 15
7 1:1 MeOH/acetate buffer 18
8 1:1 Chloroform/acetate buffer 0% of 6a 27% of
aReaction conditions: 5a (1equiv) and 2a (10equiv.) was stirred with laccase (4000U/1g substrate) in solvent at 3 °C for 2 hours and then stirred at room temperature for another 22 hours. bIsolated yield.
After developing the optimum reaction conditions, the reaction of a variety of
catechol substrates with diene 2a were examined by using the procedure for the synthesis
of o-naphthoquinones in the experimental section and these results are summarized in
Table 12. The results show that the reaction depended on the reactivity of the in situ-
generated o-quinones. The very high reactivity quinones, such as 3-methoxy-1,2-
benzoquinone and 4-chloro-1,2-benzoquinone, which have rich electron donating group
(OMe) and strong electron withdrawing group (Cl), respectively, did not provide good
yields of the o-naphthoquinone product (entries 4 and 5). These quinones preferently
OHHO
174
underwent dimerization and polymerization. For example, in-situ synthesis 3-methoxy-
1,2-benzoquinone by laccase from the corresponding hydroquinone yielded 32% of the
undesired product, which was generated by the decarbonylation and oxidation of the
dimerization intermediate, and only 11% of naphthoquinone product. Besides the
reactivity of the in situ-generated quinones, steric factor also affected the formation of the
product. Quinones with bulky groups provided very low yield of the products such as 4-
tert-butyl-1,2-benzoquinone yielded only 14% product for 4 day reaction, and 3,5-di-tert-
butyl-1,2-benzoquinone gave no product but 97% of it remaining in the reaction solution
(entries 6 and 7). From Table 12, the in-situ generated o-quinones with moderate
reactivity clearly exhibited higher yields of the o-naphthoquinone adduct (entries 1-3),
and 4-methyl-1,2-benzoquinone provided the highest yield (57%) in this reaction system
(entry 2).
175
Table 12. The study of laccase-catalyzed reaction of 2a with a variety of catechol substrates in aqueous medium
Entry Catechol Product Yielda (%)
1
6a 47
2
6b 57
3
6a 28
4
6c
11 and 32% of
5
- no product formed
6b
6d
14 and 15% of
7
-
no product formed 97% of quinone
aIsolated yield; b96 hour reaction.
OH
OH
Laccase
0.1M acetate buff er pH 4.53 oC - RT, 24 hours
O
O
2a
R1 R1
1:10
6
OHOH
Cl
OHOH
OHOH
CH3
OHOH
CHO
OHOHH3CO
OHOH
O
O
OO
H3CO
OCH3
OHOH
OO
O
O
O
O
CH3
O
O
O
O
H3CO
O
O
176
The versatility of this system for a variety of dienes was investigated by using 4-
methylcatechol as starting material to generate 4-methyl-1,2-benzoquinone in situ. Table
13 demonstrates that many dienes can be used to react with 4-methyl-1,2-benzoquinone
to generate o-naphthoquinone products in moderate to high yield. Optimal results were
achieved when 1-methoxy-1,3-butadiene and 1-acetoxy-1,3-butadiene were used as diene
reagent (entries 4 and 5). Both dienes provided very high yields of the product, and only
2 equivalence of 1-acetoxy-1,3-butadiene was needed. This high yielding reaction can be
attributed to the elimination of the methoxy or acetoxy group that ‘pushed’ the reaction
forward to the product. The proposed mechanism of the elimination of the methoxy or
acetoxy is illustrated in Figure 107. During the Diels-Alder reaction step, the steric effect
of the substituent make the reaction occurred only at the less substituent side.
177
Table 13. The study of laccase-catalyzed reaction of 4-methylcatechol with a variety of dienes in aqueous medium
Entry Diene Product Yielda (%)
1
6b 57
2
6e 71
3
6f 10
4
6g 77
5b
6g 76
6
- no product formed
aIsolated yield; bOnly 2 equivalence of 1-acetoxy-1,3-butadiene was used.
OH
OH
R4
Laccase
0.1M acetate buffer pH 4.53 oC - RT, 24 hours
O
O R3
R4
1 : 10
R3
CH3 CH3
R2
R5R5
R2
6
OCH3
OCH3
OCH3
OOCH3
O
O
CH3
O
O
CH3
O
O OCH3
CH3
OCH3
O
O
CH3
O
O
CH3
178
Figure 107. The proposed mechanism for the elimination of methoxy or acetoxy from the reaction of 4-methyl-1,2-benzoquinone and 1-methoxy-1,3-butadiene or 1-acetoxy-1,3-butadiene in the presence of laccase in aqueous medium.
In this study, we also conducted p-naphthoquinone synthesis by using a variety of
1,4-benzohydroquinones as a substrate for laccase to generate 1,4-benzoquinone in situ.
As the result of o-quinone reaction above, the reactive 1-acetoxy-1,3-butadiene was
chosen for this study. However, we found that the reaction of these less reactive p-
benzoquinones gave very low yield of the desired product at low temperature. Therefore,
the reaction was conducted at 55 °C for p-naphthoquinone synthesis, and 1 equivalence
of 1,4-benzohydroquinone and 2 equivalence of diene were used (Table 14). The
procedure for p-naphthoquinone synthesis is summarized in the experimental section.
The results in Table 14 show that this reaction system can be used for a one-pot synthesis
of p-naphthoquinones in excellent overall yield.
OH
CH3
HO
O
CH3
O
+
O
O
CH3
Laccase
OROR
O
O
CH3
OR
H OH
O
O
CH3
ROH
O
O
CH3
Diels-Alder Reaction
Dehydrogenation
- ROH
R = Me or COCH3
H
179
Table 14. The study of laccase-catalyzed reaction of 1-acetoxy-1,3-butadiene with a variety of 1,4-benzohydroquinone in aqueous medium at 55 oC
Entry R1 Product Yielda (%) 1 H 7a 67 2 CH3 7b 75 3 OCH3 7c 81 4 Br 7d 67 5 Cl 7e 69 aIsolated yield.
OH
Laccase
0.1M acetate buffer pH 4.555 oC, 24 hours
O
1:2
R1
OH
O CH3
O
O
R1
7
180
5.4 Conclusions
In summary, an efficient green chemistry synthesis of o-naphthoquinone using
laccase as an oxidant in aqueous medium was developed. In this reaction, laccase was
used to oxidize o-diphenols to generate o-quinones in situ which further underwent Diels-
Alder reaction and oxidation to form o-napthoquinone product. Due to the high reactivity
of the in situ-generated o-quinones, the reactions have to conduct at low temperature (3
oC to room temperature) to retard the side reactions, dimerization and polymerization.
This reaction system can yield o-naphthoquinones up to 77% depending on the exact
structure of the starting hydroquinone and diene. The reactions of 1-methoxy-1,3-
butadiene and 1-acetoxy-1,3-butadiene provided very high yields of the product. This
high yielding reaction can be attributed to the elimination of the methoxy or acetoxy
group that ‘pushed’ the reaction forward to the product. In addition, this study also shows
that the reaction of the reactive 1-acetoxy-1,3-butadiene and 1,4-hydroquinones catalyzed
by laccase provided the yield of the corresponding p-naphthoquinones up to 80%.
181
CHAPTER 6
CASCADE SYNTHESIS OF BENZOFURAN DERIVATIVES VIA
LACCASE OXIDATION−MICHAEL ADDITIONiii
6.1 Introduction
The provious Chapters reported the green cascade syntheses of p- and o-
naphthoquinone derivatives via Diels-Alder reaction catalyzed by laccase in aqueous
medium [379,380]. These reactions provided the satified results for the sythesis of
corresponding naphthoquinone products. To demonstrate another synthetic research
capability of laccase, herein, this Chapter presents the first laccase-catalyzed carbon-
carbon bond formation via oxidation-Michael addition for the cascade synthesis of
benzofuran derivatives. Benzofurans have attracted much attention due to their broad
spectrum of pharmacological activities [381-386] such as, anticancer, antimicrobial,
antioxidant, and anti-HIV-1 activities. Therefore, the syntheses of benzofuran derivatives
have been extensively investigated [387-391]. Most of these synthetic methods involve
the formation of an annellated furan ring by the intramolecular cyclization of benzene
iii This manuscript was published in Tetrahedron, 2007, 63, 10958-10962. It is entitled as “Cascade synthesis of benzofuran derivatives via laccase oxidation-Michael addition”. The other authors are Dr. Leslie Gelbaum and Dr. Arthur J. Ragauskas from the School of Chemistry and Biochemistry at the Georgia Institute of Technology. This chapter is reproduced with the kind permission of from [Tetrahedron]. Copyright © 2007 Elsevier Ltd.
182
derivatives. These procedures involve either multi-steps, rigorous reaction conditions, or
expensive reagents. Recently, Nematollahi et al. [392-395] and Bu et al. [396] reported
the one-pot synthesis of polyhydroxylated benzofurans via the oxidation of catechols by
an electrochemical method or sodium iodate, respectively, in the presence of 1,3-
dicarbonyl compounds. Nevertheless, using biocatalysis in the preparation of
polyhydroxylated benzofurans has never been reported. This study reports the first study
at accomplishing this synthesis via a biocatalyst.
In this procedure, ortho-quinones, generated in situ from the oxidation of
catechols by laccase, underwent the Michael addition reaction with 1,3-dicarbonyl
compounds, and then, underwent intramolecular cyclization to benzofuran derivatives
(see Figure 108). In addition, this study investigated the reaction system in the presence
of either Lewis acid or Lewis base to improve reaction condition, and documented the
recyclability of the catalytic system.
183
6.2 Experimental Section
6.2.1 General Information
All chemicals were obtained from Aldrich and used as received without further
purification. Laccase (EC 1.10.3.2) from Trametes Villosa was donated by Novo Nordisk
Biochem, North Carolina. 1H and 13C NMR spectra were recorded on a Bruker-400
spectrometer operating at 400 MHz for 1H and 100 MHz for 13C. For HMBC correlations,
the experiment was operated in a Bruker-DRX 500 spectrometer. Column
chromatography was performed on Combiflash Companion instrument (Teledyne Isco
company) using RediSep normal-phase flash columns. TLC was performed on aluminum
sheets precoated with silica gel 60 F254 (EMD Chemicals). Mass spectra were carried
out in The Georgia Institute of Technology Bioanalytical Mass Spectrometry Facility.
6.2.2 Enzyme Assay
Laccase activity measurement is described in Chapter 3 (Experimental Materials
and Procedures).
6.2.3 General Procedure of the Synthesis of Benzofuran Derivatives via Laccase-
Oxidation-Michael Addition.
The detail of the reaction procedure is described in Chapter 3 (Experimental
Materials and Procedures).
184
6.2.4 Product Characterization
Products 9a [394], 9b [396], and 9c [394] are known compounds, and their 1H-
NMR and 13C NMR data are consistent with those in literature. Structure 9d is, to the best
of our knowledge, new compounds and its NMR spectra are shown in Appendix A.3.
3-Acetyl-5,6-dihydroxy-2-methylbenzofuran (9a)
White solid; mp. 238-239 ˚C (lit. [394], 236-238 ˚C); 1H NMR (DMSO-d6, 400 MHz) δ
2.51(s, 3H, CH3), 2.67 (s, 3H, CH3), 6.92 (s, 1H, Ar-H), 7.35 (s, 1H, Ar-H), 9.03 (br s,
1H, OH), 9.10 (br s, 1H, OH); 13C NMR (DMSO-d6, 100 MHz) δ 15.2, 30.7, 97.7, 106.3,
117.1, 117.2, 143.4, 144.1, 146.9, 160.6, 193.8; MS (EI) m/z 206(M+, 92%), 191 (100),
163 (36), 135 (14), 95 (6), 89 (4), 63 (4), 53 (3), 43 (19); HRMS (EI)
206.05838(C11H10O4 requires 206.05791).
Ethyl-5,6-dihydroxy-2-methyl-3-benzofuran carboxylate (9b)
White solid; mp. 180-182 ˚C (lit. [396,397], 180-182 ˚C); 1H NMR (DMSO-d6, 400
MHz) δ 1.34 (t, J = 7 Hz, 3H, CH3), 2.62 ( s, 3H, CH3), 4.27 (q, J = 7 Hz, 2H, CH2),
6.91(s, 1H, Ar-H), 7.22 (s, 1H, Ar-H), 9.03 ( br s, 1H, OH), 9.11 (br s, 1H, OH); 13C
NMR (DMSO-d6, 100 MHz) δ 14.1, 14.3, 59.8, 97.8, 105.9, 108.1, 116.9, 143.4, 144.2,
147.1, 161.1, 163.8; MS (EI) m/z 236(M+, 92%), 207 (100), 191 (33), 93 (4), 43 (6);
HRMS (EI) 236.07061 (C12H12O5 requires 236.06847).
3-Acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c)
White solid; mp. 218-220 ˚C (lit. [394], 217-219 ˚C); 1H NMR (DMSO-d6, 400 MHz) δ
2.24 (s, 3H, CH3), 2.49 (s, 3H, CH3), 2.68 (s, 3H, CH3), 7.22 (s, 1H, Ar-H), 8.41 (br s,
1H, OH), 9.28 (br s, 1H, OH); 13C NMR (DMSO-d6, 100 MHz) δ 8.9, 15.3, 30.7, 103.2,
185
107.1, 116.2, 117.4, 141.8, 143.1, 146.5, 160.5, 193.9; MS (EI) m/z 220(M+, 85%), 205
(100), 177(21), 149 (4), 102 (5), 43 (13); HRMS (EI) 220.07490 (C12H12O4 requires
220.07356).
Ethyl-5,6-dihydroxy-2,7-dimethyl-3-benzofuran carboxylate (9d).
White-yellow solid; mp. 183-185 ˚C; 1H NMR (DMSO-d6, 400 MHz) δ 1.35 (t, J = 7 Hz,
3H, CH3), 2.25 (s, 3H, CH3), 2.66 (s, 3H, CH3), 4.29 (q, J = 7 Hz, 2H, CH2), 7.13 (s, 1H,
Ar-H), 8.42 (s, 1H, OH), 9.31 (s, 1H, OH); 13C NMR (DMSO-d6, 100 MHz) δ 8.9, 14.2,
14.2, 59.8, 102.8, 107.2, 108.3, 115.9, 141.9, 143.0, 146.7, 161.0, 163.9; MS (EI) m/z
250(M+, 98%), 221 (100), 176(11), 93 (4), 43 (7); HRMS (EI) 250.08453(C13H14O5
requires 250.08412).
186
6.3 Results and Discussion
6.3.1 Preliminary Study and the Effect of pH on the Reaction System
In a preliminary study, the reaction of 3-methylcatechol (5b) and acetylacetone
(8a) in the presence of laccase was investigated. The reaction was carried out under air at
room temperature (23 ˚C) in the aqueous buffer solution for 4 hours. This reaction system
was chosen to be a model reaction for this study because the product, 3-acetyl-5,6-
dihydroxy-2,7-dimethylbenzofuran (9c), gradually precipitated during the reaction and
was easy to recover by filtration after the reaction.
The effect of pH on this reaction system was initially studied. As summarized in
Table 15, the optimal yields of 9c were achieved when the reaction was conducted at pH
7.0. At pH 4.5, no product formed because this low pH was not basic enough to
deprotonate alpha-proton from acetylacetone to facilitate the Michael addition reaction
with the in situ generated o-quinone. At a higher pH value of 8.0, only a small yield of 9c
was received due to laccase activity which was dramatically decreased at this pH
[199,366]. Therefore, only a small amount of starting catechol was oxidized and reacted
subsequently with acetylacetone. Moreover, the ratio of 5b and 8a also affected the yield
of 9c. The result shows that the yield of 9c increased when using 5b and 8a in 1:2 ratio
(entry 2).
187
Table 15. The effect of pH on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a)
Entry Solvent/ pH 5b:8a (equiv) Yielda of 9c (%)
1 0.1 M Phosphate buffer pH 7.0 1:1 46
2 0.1 M Phosphate buffer pH 7.0 1:2 64
3 0.1 M Acetate buffer pH 4.5 1:2 0
4 0.1 M Phosphate buffer pH 8.0 1:2 6 aIsolated yield.
6.3.2 The Effect of Lewis Bases on the Reaction System
After this preliminary study, the next phase was to improve the yield of the
product by enhancing Michael-addition step. Traditionally, Michael reactions are
catalyzed by strong bases such as alkali metal, alkoxides or hydroxides [398]. However,
these strongly basic conditions can lead to a number of side- and subsequent reactions,
and especially for this reaction system, the in situ-generated o-quinones are easily
decomposed in the presence of hydroxides [376]. Recently, Xia et al. [399] reported the
use of a Lewis base to catalyze Michael addition of azide ion to cyclic enones in water.
Herein, adding Lewis base to the catalyzed Michael addition step was investigated. Table
16 reveals that the best yield of 9c was obtained when using pyridine as Lewis base in
phosphate buffer pH 7.0, and the ratio of 5b:8a: pyridine was 1:2:1. While the use of
OH
OHO O O OH
OH
O
+LaccaseSolventRT, 4 h
5b 8a 9c
188
stronger Lewis acid such as 4-dimethylaminopyridine (DMAP) and 1,4-
diazabicyclo[2.2.2]octane (DABCO) provided only a low yield of the product. Although
the use of pyridine gave the best result for this reaction system, the yield of the product
(54%, entry 3) was still much lower than the yield of the product (64%, Table 15, entry 2)
accomplished without pyridine. According to these results, adding basic reagents into this
reaction did not enhance the reaction efficiency, especially, when a strong base was used.
Table 16. The effect of Lewis bases on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a)
Entry Lewis bases Solvent 5b: 8a: Lewis base (equiv)
Yielda of 9c (%)
1 Pyridine Water 1: 2: 0.5 33
2 Pyridine 0.1 M Phosphate buffer pH 7.0
1: 2: 0.5 40
3 Pyridine 0.1 M Phosphate buffer pH 7.0
1: 2: 1 54
4 DMAP 0.1 M Phosphate buffer pH 7.0
1: 2: 1 9
5 DABCO 0.1 M Phosphate buffer pH 7.0
1: 2: 1 13
aIsolated yield.
OH
OHO O O OH
OH
O
+Laccase
SolventRT, 4 h
5b 8a 9c
Lewis base
189
6.3.3 The Effect of Lewis Acids on the Reaction System
In order to circumvent the alkaline conditions above, we decided to investigate
the reaction in the presence of a Lewis acid as an alternative method. Lewis acid-
catalyzed Michael reactions have been developed, allowing the reaction to be carried out
under milder conditions with high efficiency [400]. Our studies focus on the use of water
as reaction medium to avoid the use of organic solvents which have become an
environmental concern. Studies by Kobayashi et al. have showed that the rare earth metal
triflates (Sc(OTf)3, Yb(OTf)3, etc.) can be used as Lewis acid catalyst in water-containing
solvents [401,402]. Therefore, we examined a variety of Lewis acids including the water-
compatible Lewis acid, Sc(OTf)3 and Yb(OTf)3, for the synthesis of 9c. The reaction was
carried out under the optimal condition in the preliminary study (Table 15, entry 2) but
Lewis acids were varied. The results of this Lewis acid study is summarized in Table 17.
The results show that the water-stable Lewis acid, Sc(OTf)3 and Yb(OTf)3 can enhance
Michael addition step for this reaction system and provided a very good yield of 9c.
Sc(OTf)3 showed better result than Yb(OTf)3. However, we have to use 0.2 equiv of
Sc(OTf)3 to obtain the highest yield of 9c (74%, entry 2) because using only 0.1 equiv of
Sc(OTf)3 did not have any effect on the reaction yield (63%, entry 1) when compared to
the reaction without Sc(OTf)3 (64%, Table 15, entry 2).
As we conducted the reaction in aqueous medium, the main drawback was the
low solubility of organic substances. To overcome this problem, a small amount of
surfactant, sodium dodecyl sulfate (SDS, 20 mol %) was used to improve the solubility,
and the result shows a small increase of product yield from 74% to 76% (Table 17, entry
190
3). This result agrees with Kobayashi’s work on the study of surfactant-aided Lewis acid
catalysis in aqueous aldol reaction [403].
Table 17. The effect of Lewis acids on the laccase-catalyzed reaction of 3-methylcatechol (5b) and acetylacetone (8a)
Entry Lewis acid 5b: 8a: Lewis acid (equiv) Yielda of 9c (%)
1 Sc(OTf)3 1: 2: 0.1 63 2 Sc(OTf)3 1: 2: 0.2 74 3 Sc(OTf)3/ SDS 1: 2: 0.2 76 4 Yb(OTf)3 1: 2: 0.2 72 5 InCl3.4H2O 1: 2: 0.2 71 6 CuCl2 1: 2: 0.2 49 aIsolated yield
6.3.4 The Synthesis of Benzofuran Derivatives
After successfully conducting the optimization experiments described above, we
chose to conduct further synthesis of benzofuran derivatives by introducing 1 mmol of
substituted catechols and 2 mmol of 1,3-dicarbonyl compounds in 0.1M phosphate buffer
(pH 7.0), in the presence of laccase, 20 mol% of Sc(OTf)3, and 20 mol% of SDS under
air at room temperature. The proposed reaction pathway of this catalytic system is
OH
OHO O O OH
OH
O
+Laccase, Lewis acid
RT, 4 h
5b 8a 9c
Phosphate buff er pH 7
191
illustrated in Figure 108, and the result of the reaction of various catechols and 1,3-
dicarbonyl compounds are summarized in Table 18.
Figure 108. Proposed mechanism of laccase/Sc(OTf)3 catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c).
O
O
(TfO)3Sc
O
O
OH
OH
Laccase (ox)Laccase (red)
O2 from Air H2O
O
O
O
O
(TfO)3Sc
O
O
O
O
(TfO)3Sc
Sc(OTf)3
O O
HO
O
O
O
HO
O
O
OHO
O
HO
HO O
O
HO
HO
Aromatization
5b
8a
9c
192
Table 18. The study of the laccase/Sc(OTf)3-catalyzed reaction of catechols and 1,3-dicarbonyl compounds for benzofuran synthesis
Entry
1 5a: R1 = R2 = H 8a: R3 = R5 = Me, R4 = H 9a (68%) 2 5a 8b: R3 = R5 = Me, R4 = Cl 9a (66%)b 3 5a 8c: R3 = Me, R4 = Cl, R5 = OEt 9b (46%)b 4 5b: R1 = Me, R2 = H 8a 9c (76%) 5 5b 8b 9c (79%)b 6 5b 8c 9d (48%)b 7 5c: R1 = OMe, R2 = H 8a - 8 5d: R1 = F, R2 = H 8a - 9 5e: R1 = H, R2 = Cl 8a 9a (9%) 10 5f: R1 = H, R2 = COOH 8a 9a (11%) aIsolated yield; bReaction time is 1 hour.
The data in Table 18 show that the reaction depends on the reactivity of the in
situ-generated o-quinones. The very reactivity quinones, such as 3-methoxy-1,2-
benzoquinone and 3-fluoro-1,2-benzoquinone, which have rich electron donating group
(OMe) or a strong electron withdrawing group (F), respectively, did not provided any
desired products (entries 7 and 8). This reactivity pattern may be caused by side reactions
of these highly reactive quinones. In contrast, the reaction of catechols, such as 3-
R1
OH
OH
R3 R5
O O O
R1
OH
OH
R3
R5
O
+Laccase,
0.1M Phosphate buffer pH 7RT, 4 hR2 R4
0.2 eq. Sc(OTf)3
0.2 eq. SDS,
R1
OHOH
5
Catechol
R2
R3 R5
O O
8R4
1,3-Dicarbonyl compound
O
R1
OH
OH
R3
R5O
9
Product (%yield)a
193
methylcatechol and catechol with laccase generated moderately reactive quinones that
gave excellent yields of the corresponding benzofuran products as shown in entries 1-6.
Moreover, the reactivity of 1,3-dicarbonyl compounds also have an effect on the reaction.
When we used 1,3-dicarbonyl compounds that had an electron withdrawing group (Cl) at
alpha-position, the reaction time was only 1 hour. The shorter reaction time caused by the
increase of alpha-proton acidity of these 1,3-dicarbonyl compounds that make it easier to
deprotonate and ready to react with in situ-generated o-quinone in the reacion solution.
Besides 3-substituted catechols, 4-substituted catechols, such as 4-chlorocatechol and
3,4-dihydroxy benzoic acid, can also be used for the synthesis of polyhydroxylated
benzofurans (entries 9 and 10). However, the yield of the product is low.
In addition, we observed that this reaction system gave only one isomer from
potential products that could occur. This could be explained by the existence of a
substituent at the C-3 position of catechols that probably causes the Michael acceptors, in
situ generated o-quinones, to be attacked by 1,3-dicarbonyl compounds only at less
hindered C-5 position to yield the observed product (see Figure 108). Most of the
products from this study are known compounds. Only product 9d is unknown. Therefore,
the structure of 9d was confirmed by the 1H NMR, 13C NMR and HMBC correlations as
shown in Table 19.
194
Table 19. 1H and 13C assignment and HMBC correlations for compound 9da
Carbon 13C (δ) 1H (δ) 1H-13C Correlations
2a 14.2 2.66, s, 3H C2, C3 4 102.8 7.13, s, 1H C3, C5, C6, C7a 8 8.9 2.25, s, 3H C6, C7, C7a 2’ 59.8 4.29, q, 2H (7) C1’, C3’ 3’ 14.2 1.35, t, 3H (7) C2’ OH (5) 9.31, s, 1H C4, C5 OH (6) 8.42, s, 1H C6, C7 aMeasure in DMSO-d6 at 125 (13C) or 500 MHz (1H, J (Hz) values in parentheses). Chemical shifts are expressed in δ(ppm). The HMBC spectrum is shown in Appendix A.3.
6.3.5 The Recyclability of the Laccase/Sc(OTf)3-Catalytic System
Next, we examined the recyclability of the two-component catalytic system,
laccase/Sc(OTf)3, for the synthesis of benzofuran 3a by the reaction of 5b and 8a in 0.10
M phosphate buffer pH 7.0 and 20 mol% SDS. Due to the product 9c precipitated during
the reaction, we can directly reuse this catalytic system after product filtration. The
results shown in Table 20 demonstrate that this catalytic system was readily recyclable
for three runs, with approximately a 10% drop of the product yield/reaction.
O OH
OH
O
O9d
2
3 45
67
8
2a
2'3' 1'
3a
7a1
195
Table 20. Recycling of the laccase/Sc(OTf)3 catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c)
Run Yielda of 9c (%)
1 76 2 62 3 51 aIsolated yield.
OH
OH
O O
O OH
OH
O
+Laccase,
0.1M Phosphate buffer pH 7rt, 4 h
0.2 eq. Sc(OTf)3
0.2 eq. SDS,
5b
8a
9c
196
6.4 Conclusions
In conclusion, this study provides an efficient green chemistry synthesis of
benzofuran derivatives from the reaction of catechols and β-dicarbonyl compounds using
a catalytic system of laccase and Sc(OTf)3 in surfactant aqueous medium. This reaction is
regioselective providing only one isomer product and the first example of a two
component catalytic system employing laccase and a lanthanide Lewis acid catalyst. The
yield of the products from reaction depended on both the reactivity of catechols and β-
dicarbonyl compounds. For this reaction system, catechols with moderate reactivity yield
benzofuran products in excellent yield. In addition, the newly developed catalytic system
could also be recycled and reused for two additional runs, with only a minor drop in
product yields.
197
CHAPTER 7
CO-CATALYTIC ENZYME SYSTEM FOR THE MICHAEL
ADDITION REACTION OF IN SITU-GENERATED
ORTHO-QUINONESiv
7.1 Introduction
In recent years, the advances in genomics and biotechnology have dramatically
broadened the availability of low-cost enzymes. In turn, this has increased the potential
application of enzymes for organic synthesis while also addressing the challenges of
green chemistry [32]. A growing field of interest in this field is the application of
enzyme-initiated domino reactions [1,113-115]. Under optimized reaction conditions it
has been shown that several biocatalytic reactions can be carried out in a single reactor
[137-146]. For example, Kroutil and his co-workers [148] recently developed the one pot,
two step, two enzyme cascade reaction for the synthesis of enantiopure epoxide. Herein,
we report on the use of two enzymes, laccase and lipase, in the domino reaction of in
situ-generated o-quinones followed by enzyme catalyzed Michael addition.
iv This manuscript was submitted to European Journal of Organic Chemistry, 2008. It is entitled as “Co-catalytic enzyme system for the Michael addition reaction of in situ-generated ortho-quinones”. The other author is Dr. Arthur J. Ragauskas from the School of Chemistry and Biochemistry at the Georgia Institute of Technology
198
Although lipases (triacylglycerol hydrolase, EC 3.1.1.3) have been known to
catalyze the hydrolysis and the synthesis of esters formed from alcohols and acids
[304,305,318], recent studies have reported the ability of lipases to catalyze Michael
addition reactions [321,322,325]. For example, Torre et al. [321] provided the initial
demonstration that lipase was able to catalyze the Michael addition of secondary amines
to acrylonitrile. This reaction is clearly different from the natural process this enzyme is
usually associated with. Berglund et al. [325] has reported the Michael addition of 1,3-
dicarbonyl compounds to α,β-unsaturated carbonyl compounds catalyzed by a C.
antarctica lipase B mutant. Moreover, Wang et al. [322] recently established that lipase
M from Mucor javanicus was able to catalyze the Michael addition reaction of
pyrimidine with a disaccharide acrylate.
According to Chapter 6, an aqueous cascade synthesis of benzofuran derivatives
from the reaction of catechols and 1,3-dicarbonyl compounds via an oxidation-Michael
addition sequence catalyzed by laccase and Sc(OTf)3/SDS was successfully developed
[404]. Depending on the exact substrates, one-pot yields of benzofurans averaged 50-
79% and in the absence of Sc(OTf)3, these yields decreased to 45-65%. Hence, the use of
an aqueous Lewis acid was critical for efficient synthesis of the desired compounds. In
regards to environmental concern, this system still produces some hazardous waste from
the metal catalyst. Therefore, the development of alternative methodologies to replace the
lanthanide metal catalyst in this synthesis is a high priority to enhance the overall green
chemistry aspect of this one-pot synthetic reaction. This Chapter presents the use of
enzyme, lipase, as an alternative catalyst in conjunction with laccase for the synthesis of
benzofuran derivatives. In addition, in this study, this lipase/laccase co-catalytic system
199
was also used to catalyze the Michael addition of in situ-generated o-quinones and
aromatic amines.
7.2 Experimental Section
7.2.1 General Information
All chemicals were used as received without further purification. Laccase (EC
1.10.3.2) from Trametes villosa was donated by Novo Nordisk Biochem, North Carolina.
Lipases were purchased from Aldrich. Unit definition of each lipase is different
depending on the method that Aldrich used to measure lipase activity. The enzymes were
kept frozen until used. 1H and 13C NMR spectra were recorded on a Bruker-400
spectrometer operating at 400 MHz for 1H and 100 MHz for 13C in d6-DMSO or CDCl3
using tetramethylsilane (TMS) as the internal standard. All reactions were monitored by
TLC. TLC was performed on aluminum sheets precoated with silica gel 60 F254 (EMD
Chemicals). Column chromatography was performed on Combiflash Companion
instrument (Teledyne Isco company) using RediSep normal-phase flash columns. Mass
spectra were carried out in The Georgia Institute of Technology Bioanalytical Mass
Spectrometry Facility.
200
7.2.2 Enzyme Assay
Laccase activity measurement is described in Chapter 3 (Experimental Materials
and Procedures).
7.2.3 General Procedure of the Synthesis of Benzofuran Derivatives Using Laccase-
Lipase Co-Catalytic System.
The detail of the reaction procedure is described in Chapter 3 (Experimental
Materials and Procedures).
7.2.4 Procedure for the Study of the Reaction of 5a and 8a (with and without
Lipase PS)
In a 250-mL round-bottom flask, 40 ml of 0.10 M phosphate buffer pH 7.0 and 5a
(2 mmol, 0.2202 g) were mixed together. Next, 200 U of laccase was added to reaction
mixture and then, 8a (0.4004 g, 410 μl, 4 mmol ) and 1848 U of lipase PS (or no lipase)
were added. The reaction was then stirred at room temperature in a flask open to the
atmosphere for 4.5 h. A 3 ml aliquot of the reaction mixture was taken every 30 minutes
during the reaction and extracted with 10 ml of EtOAc. The organic phase was then dried
over MgSO4, filtered and concentrated under reduced pressure. The resulting crude
product was submitted to quantitative 1H NMR analysis to measure the formation of
product 9a using 0.5 ml of 0.20 M 1,3,5-trioxane in d6-DMSO as internal standard.
Figure 109 illustrates 1H-NMR spectra of the crude mixture that show the formation of
product 9a during the reaction. Ar-H peaks of 9a are used to calculate the yield of 9a.
201
Figure 109. 1H-NMR of crude mixture from the laccase-catalyzed the reaction of 5a and 8a with and without lipase. These spectra demonstrate the formation of 9a and the decrease of starting material 5a during the reaction.
7.2.5 General Procedure for the Reaction of Catechols and Anilines Catalyzed by
Laccase-Lipase Co-Catalytic System.
The detail of the reaction procedure is described in Chapter 3 (Experimental
Materials and Procedures).
202
7.2.6 Product Characterization
Compond 9a [394], 9b [395], 9c [394], 9d [404], and 11a [405] are known
compounds and our 1H and 13C NMR data are consistent with those in the literature.
Structure 11b, 11c, and 11d are, to the best of our knowledge, new compounds. 1H and
13C assignments and HMBC correlation for compound 11b, 11c, and 11d are summarized
in Table 21. These NMR spectra are shown in Appendix A.4.
Compound 11a
Red solid: Yield: 87 mg (30%). m.p. 193-195 ºC. 1H NMR (400 MHz, CDCl3): δ = 8.59
(br s, 1H, OH), 7.55 (br s, 1H, OH), 7.42 (t, J = 9 Hz, 4 H, 4 × CH arom.), 7.22 (t, J = 9
Hz, 2 H, 2 × CH arom.), 7.12 (br s, 4 H, 4 × CH arom.), 6.10 (s, 2 H, 2 × CH) ppm. 13C
NMR (100 MHz, CDCl3): δ = 96.0, 121.9, 125.7, 129.4 ppm. MS (EI): m/z = 290 (M+,
70%), 261 (26), 144 (15), 77 (23), 51 (8). HRMS (EI): calcd. for C18H14N2O2 290.1055;
found 290.1038.
Compound 11b
Red solid. Yield: 129.5 mg (37%). m.p. 161-162 ºC. 1H NMR (400 MHz, CDCl3): δ =
8.50 (br s, 1 H, OH), 7.56 (br s, 1 H, OH), 7.08 (d, J = 7 Hz, 4 H, 4 × CH arom.), 6.94 (d,
J = 8 Hz, 4 H, 4 × CH arom.), 6.07 (s, 2 H, 2 × CH), 3.84 (s, 6 H, 2 × OCH3) ppm. 13C
NMR (100 MHz, CDCl3): δ = 157.4, 151.9, 135.3, 123.9, 114.2, 95.1, 55.1 ppm. IR
(KBr): υmax = 3293 (s), 3246 (s), 3040 (w), 2834 (w), 1739 (w), 1654 (w), 1606 (s), 1580
(s), 1525 (s), 1511 (s), 1411 (s), 1330 (m), 1286 (m), 1244 (s), 1217 (s), 1199 (s), 1173
(m), 1033 (m), 840 (m)cm-1. MS (EI): m/z = 350 (M+, 86%), 319 (100), 291 (12), 174
(15), 146 (12), 92 (7), 77 (9). HRMS (EI): calcd. for C20H18N2O4 350.1266; found
350.1247.
203
Compound 11c
Red solid. Yield: 182.6 mg (51%). m.p. 219-221 ºC. 1H NMR (400 MHz, DMSO-d6): δ =
9.24 (br s, 2 H, 2 × OH), 7.44 (d, J = 7 Hz, 4 H, 4 × CH arom.), 7.19 (br s, 4 H, 4 × CH
arom.), 5.81 (s, 2 H, 2 × CH) ppm. 13C NMR (100 MHz, DMSO-d6): δ = 152.0, 142.6,
129.1, 128.9, 123.7, 97.5 ppm. IR (KBr): νmax = 3298 (s), 3031 (w), 1736 (w), 1660 (w),
1606 (m), 1573 (s), 1536 (s), 1493 (s), 1480 (s), 1415 (s), 1334 (s), 1221 (s), 1188 (s),
1087 (m), 1007 (m), 830 (m) cm-1. MS (EI): m/z = 358 (M+, 42%), 323 (80), 288 (8), 178
(18), 144 (15), 127 (100), 84 (57), 65 (18), 49 (75). HRMS (EI): calcd. for
C18H12N2O2Cl2 358.0275; found 358.0266.
Compound 11d
Red solid. Yield: 159 mg (50%). m.p. 194-196 ºC. 1H NMR (400 MHz, CDCl3): δ = 8.55
(br s, 1 H, OH), 7.55 (br s, 1 H, OH), 7.20 (d, J = 7 Hz, 4 H, 4 × CH arom.), 7.02 (br s, 4
H, 4 × CH arom.), 6.09 (s, 2 H, 2 × CH), 2.37 (s, 6 H, 2 × CH3) ppm. 13C NMR (100
MHz, CDCl3): δ = 152.2, 135.5, 129.9, 122.0, 95.7, 20.9 ppm. IR (KBr): νmax = 3297 (s),
3031 (w), 2917 (w), 1739 (m), 1663 (w), 1600 (s), 1572 (s), 1533 (s), 1511 (s), 1488 (s),
1413 (s), 1337 (s), 1219 (s), 1189 (s), 1153 (s), 897 (m), 814 (m), 732 (m) cm-1. MS (EI):
m/z = 318 (M+, 42%), 303 (100), 275 (15), 158 (13), 130 (8), 91 (14), 65 (8), 49 (11).
HRMS (EI): calcd. for C20H18N2O2 318.1368; found 318.1348.
204
Table 21. 1H and 13C assignments and HMBC correlation for compound 11b, 11c, and 11d.a
Compound 11b Carbonb 13C (δ) 1H (δ) 1H-13C correlation 2, 2’ 95.1 6.07, s, 2H C3, C3’ 5, 5’, 9, 9’ 123.9 7.08, d, 4H (7) C4, C4’, C6, C6’, C7, C7’, C8, C8’ 6, 6’, 8, 8’ 114.2 6.94, d, 4H (8) C4, C4’, C5, C5’, C7, C7’, C9, C9’ 10, 10’ 55.1 3.84, s, 6H C7, C7’ OH (1, 1’) 7.56, br s, 1H
8.50, br s, 1H
Compound 11c Carbonc 13C (δ) 1H (δ) 1H-13C correlation 2, 2’ 97.5 5.81, s, 2H C3, C3’ 5, 5’, 9, 9’ 129.1 7.44, d, 4H (7) C4, C4’, C6, C6’, C8, C8’ 6, 6’, 8, 8’ 123.7 7.19, br s, 4H C5, C5’, C7, C7’, C9, C9’ OH (1, 1’) 9.24, br s, 2H
Compound 11d Carbond 13C (δ) 1H (δ) 1H-13C correlation 2, 2’ 95.7 6.09, s, 2H C3, C3’ 5, 5’, 9, 9’ 122.0 7.02, br s, 4H C6, C6’, C7, C7’, C8, C8’ 6, 6’, 8, 8’ 129.9 7.20, d, 4H (7) C5, C5’, C9, C9’, C10, C10’ 10, 10’ 20.9 2.37, s, 6H C6, C6’, C7, C7’, C8, C8’ OH (1, 1’) 7.55, br s, 1H
8.55, br s, 1H
aMeasure in CDCl3 or DMSO-d6 at 100 MHz (13C) or 400 MHz (1H, J (Hz) values in parentheses). Chemical shifts are express in δ (ppm); bCompound 11b: 13C (δ) of C-3/3’, C-4/4’ and C7/7’ = 151.9, 135.3, and 157.4 ppm; cCompound 11c: 13C (δ) of C-3/3’, C-4/4’ and C7/7’ = 152.0, 142.6, and 128.9 ppm; dCompound 11d: 13C (δ) of C-3/3’ and C7/7’ = 152.2 and 135.5 ppm.
OHHO
NN RR
11
11b: R = OCH311c: R = Cl11d: R = CH3
1
2
3
1'
2'
3'
4 4'
5 5'6 6'
7
8 9 8'9'
7'
10 10'
205
7.3 Results and Discussion
7.3.1 Laccase-Lipase Co-Catalytic System for the Reaction of Catechols and 1,3-
Dicarbonyl Compounds
In this study, laccase first catalyzed the oxidation of catechols to the
corresponding o-quinones which were reacted in-situ with 1,3-dicarbonyl compounds via
a Michael addition reaction. The Michael addition step was catalyzed by lipase and the
resulting addition product undergoes a subsequent intramolecular cyclization to form
benzofuran derivative products (see Figure 110).
Figure 110. Proposed reaction pathway of laccase/lipase catalytic system for the synthesis of compound 9a.
OH
OH
5a
O
OLaccase
O O
Lipase
8a
OH
O
OO
O
O
OHO
O
O
O
HO
O
O
OH
HO
Aromatization
9a
O2 H2O
-2H+
H
206
In our initial studies, the reaction of catechol (5a) and acetylacetone (8a) in the
presence of laccase and lipase from Candida rugosa (lipase CR, 60,000U/mg) was
investigated. The reaction was carried out under atmospheric conditions at room
temperature (23 ˚C) in an aqueous buffered solution for 4 hours. We found that the
optimal yield of the product (9a) of 60% was achieved when conducting the reaction of
5a and 8a in 1:2 molar ratio at pH 7.0, and using 100 U of laccase and 10 mg (600,000
U) of lipase CR per 1 mmol of 5a. Because of the high activity of in situ-generated
quinone, some side products (e.g. from the polymerization of the quinone) were also
observed but in this study we did not separate and indentify them. For the control reaction
when no laccase and lipase was added, no product was formed. When this reaction was
preformed using only lipase no product was formed, and in the presence of laccase only,
the product 9a was formed in only 33% yield.
After this preliminary study, the next phase was to examine a variety of lipases
for this reaction system. The esterases studied included lipase CR (60,000U/mg), lipase
from Pseudomonas cepacia (lipase PS, 46.2U/mg), and lipase B Candida antarctica
(CALB, 10.8U/mg). The activity of these lipases was measured by Aldrich methods
which are different for each lipase. The catalytic properties of these lipases were
investigated by reacting 8a with catechols, 5a and 3-methylcatechol (5b), in the presence
of laccase, as summarized in Table 22. This study established that the optimal amount of
each lipase to provide the highest yield of the product was different. The optimal amount
of lipase CR, lipase PS and CALB for the reaction conditions used was 600,000 U, 924
U, and 54 U per 1 mmol of catechol, respectively. The data in Table 22 shows that the
yield of the product usually increased when lipase was added to the reaction. Lipase PS
207
and lipase CR gave a high yield of the products for both reactions while CALB was good
only for reaction 2. In addition, lipase PS activity used in the reaction was much less
than of lipase CR. Therefore, lipase PS was chosen for further study. In order to verify
whether the lipase reaction is indeed catalyzed by the active site of lipase PS and not by
the protein, the reactions using inactivated lipase PS were conducted. The results in Table
22 show that the inactivated lipase PS showed no catalytic activity for these reactions.
Table 22. Reaction of catechol (5a) and acetylacetone (8a) in the presence of laccase with a variety of lipases.
Lipase Yield (%)a
No lipase 33 Inactivated lipase from Pseudomonas cepacia 31 Lipase from Candida rugosa (Lipase CR) 60 Lipase from Pseudomonas cepacia (Lipase PS) 58 Lipase B Candida antarctica (CALB) 41
aIsolated yield.
OH
OH O O O OH
OH
O
+Laccase, Lipase
Phosphate Buffer pH 7.0rt, 4h
5a 8a 9a
(1)
208
Table 22. (Continued)
Lipase Yield (%)a
No Lipase 53 Inactivated lipase from Pseudomonas cepacia 50 Lipase from Candida rugosa (Lipase CR) 56 Lipase from Pseudomonas cepacia (Lipase PS) 60 Lipase B Candida antarctica (CALB) 62
aIsolated yield.
To further define the catalytic benefits of lipase PS, the reaction of 5a and 8a in
the presence of laccase with and without lipase PS were carried out. Sample aliquots
were taken every 30 minutes during the reaction and a quantitative analysis of product 9a
was measured by 1H-NMR spectroscopy using 1,3,5-trioxane as an internal standard. The
calculated yield of the product 9a is higher than the isolated yield shown in Table 22 in
both cases (with and without lipase PS). This can be explained by the losing of product
yield during the isolation process. However, in the end of reaction, the yield difference
between the reaction with and without lipase is about the same which is approximately
25%. Figure 111 shows that in the beginning of the reaction, the rate and yield of 9a from
both reactions were almost the same. This can be explained by the predominance of
laccase-catalyzed oxidation of catechol at the beginning of the reaction. At this stage,
catechol was gradually oxidized by laccase which led to a low concentration of o-
OH
OH O O O OH
OH
O
+Laccase, Lipase
Phosphate Buffer pH 7.0rt, 4h
5b 8a 9c
(2)
209
quinone. After 2 hours of the reaction (when the concentration of laccase-generated
quinone was high enough), the reaction with lipase PS was predominant and provided a
higher rate of the reaction and higher yield of the product than the reaction without lipase
PS. Therefore, lipase PS can enhance the overall yield for this reaction system.
0
10
20
30
40
50
60
70
80
0 0.5 1 1.5 2 2.5 3 3.5 4 4.5
Yield of 9a (%
)
Time (hours)
no lipase PS
with lipase PS
Figure 111. The formation of compound 9a from the reaction of 5a and 8a in the presence of laccase. The percent yield of 9a was measured by 1H-NMR spectroscopy.
Following these optimization studies, we evaluated the breadth of this laccase-
lipase co-catalytic system for the synthesis of benzofuran derivatives using a variety of
catechols and 1,3 dicarbonyl compounds. The results summarized in Table 23 clearly
suggest that the inactivated lipase has no catalytic effect on the reactions. In addition, the
reactivity of the 1,3-dicarbonyl compound employed also has an effect on efficiency of
210
this two-enzyme system. When we used 1,3-dicarbonyl compounds that had an electron
withdrawing group (Cl) at the alpha-position, the reaction was complete in 1.5-2 hours.
The shorter reaction time was ascribed to the increased acidity of the alpha-proton of
these substituted 1,3-dicarbonyl compounds. The proposed mechanism of the elimination
of Cl atom is illustrated in Figure 112. Besides 3-substituted catechols, 4-substituted
catechols, such as 4- chlorocatechol, can also be used for the synthesis of
polyhydroxylated benzofurans (entry 11). However, the yield of the product is low. In
addition, we observed that this reaction system gave only one isomer form of the possible
benzofuran products.
Next, we examined the recyclability of this two-enzyme catalytic system for the
synthesis of benzofuran 9c. The product 9c is relatively insoluble in the aqueous reaction
mixture and readily precipitates out of solution. Simple filtration of the product mixture
facilitates reuse of the lipase/laccase reaction system. The results shown in Table 24
demonstrate that this catalytic system can be reused for a second reaction, but for the
third treatment only a low yield of the product was formed. The decrease of product yield
after the third experiment resulted from the presence of laccase inhibitor, Cl-, in the
reaction mixture that led to the decrase of laccase activity [198].
211
Table 23. The study of the laccase/lipase catalyzed reaction of catechols and 1,3-dicarbonyl compounds in aqueous medium
Entry
1 5a: R1 = R2 = H 8a: R3 = R5 = Me, R4 = H 9a (58%) 9a (31%)b 2 5a 8b: R3 = R5 = Me, R4 = Cl 9a (51%)c 9a (40%)b 3 5a 8c: R3 = Me, R4 = H, R5 = OEt 9b (11%) 4 5a 8d: R3 = Me, R4 = Cl, R5 = OEt 9b (53%)c 9b (26%)b 5 5b: R1 = Me, R2 = H 8a 9c (60%) 9c (50%)b 6 5b 8b 9c (72%)c 9c (52%)b 7 5b 8c 9d (13%) 8 5b 8d 9d (66%)c 9d (39%)b 9 5c: R1 = OMe, R2 = H 8a - 10 5d: R1 = F, R2 = H 8a - 11 5e: R1 = H, R2 = Cl 8a 9a (8%) aIsolated yield; bIsolated yield from the reaction using inactivated lipase PS; cReaction time is 1.5-2 hours.
R1
OH
OH
R3 R5
O O O
R1
OH
OH
R3
R5O
+Laccase, Lipase PS
0.1M Phosphate buffer pH 7rt, 4 hR2 R4
R1
OHOH
5
Catechol
R2
R3 R5
O O
8R4
1,3-Dicarbonyl compound
O
R1
OH
OH
R3
R5O
9
Product (%yield)a
212
Figure 112. The proposed mechanism of the elimination of Cl atom from the laccase/lipase catalyzed reaction of catechol and 8b in aqueous medium. Table 24. Recycling of the laccase/lipase co-catalytic system for the synthesis of 3-acetyl-5,6-dihydroxy-2,7-dimethylbenzofuran (9c)
Run Yielda of 9c (%) 1 72 2 62 3 5
aIsolated Yield.
OH
OHO O
O OH
OH
O
+Laccase,
0.1M Phosphate buffer pH 7rt, 1.5 h
Lipase PS
5b 8b 9c
Cl
OH
OH
5a
O
OLaccase
O O
Lipase8b
OH
O
OO
OH
O
OHO
O
O
OH
HO
O
O
OH
HO
9a
Cl
Cl
H
Michael addition
OH
O
OO
H
HO
H
O2 H2O
213
7.3.2 Laccase-Lipase Co-Catalytic System for the Reaction of Catechols and
Anilines
Next, we explored the feasibility of the laccase-lipase co-catalytic system for the
reaction between catechol and aromatic amines, anilines. Lalk and his co-worker have
demonstrated the ability to synthesize aminoquinones by laccase initiated oxidation of p-
hydroxyquinones followed by Michael addition of primary aromatic amines in a good to
excellent yields [11]. In contrast, herein, this nuclear animation reaction with the reactive
1,2-catechols was reported to yield the corresponding products less than 35%. In the
presence of lipase, we had hypothesized that thus enzyme could catalyze the Michael
addition step of the reaction between the laccase-generated o-quinone and anilines
thereby improving the overall yields. We first conducted the reaction of catechol (5a) and
aniline (10) in the presence of laccase, with or without lipase PS, in phosphate buffer pH
7.0 at room temperature for 3.5 hours. The ratio of catechol and aniline was 1 to 2, and
100U of laccase and 924U of lipase per 1 mmol of catechol were used. An insoluble red
color product precipitated out of solution during the reaction. Therefore, the product was
readily collected by filtration completion of reaction. The results show that the yield of
the reaction with lipase PS increased by ~30% when compare to the yield of the reaction
without lipase PS. Next, the amount of lipase PS used in the reaction was increased from
924U to 1848U per 1 mmol of catechol to study the effect of lipase dose on the reaction
system. This result suggests that the increase of lipase dose did not provide a significant
improvement for this reaction system (Table 25). Characterization of the product by
NMR and mass spectrum indicated that the product was composed of a 1:2 ratio of 1,2-
benzoquinone and aniline (M+/Z = 290). Moreover, the product was not a quinone
214
structure because the carbonyl carbon signal was not observed in 13C-NMR spectrum.
The proposed reaction pathway for this reaction and the structure of product (11a) was
illustrated in Figure 113. Compound 11a are known compounds and our 1H and 13C
NMR data are consistent with those in the literature [405]. In our and literature’s 13C-
NMR spectrum, peak of carbon that connect to nitrogen atom is not observed. This may
be due to the effect of nitrogen atom that broaden the peak and make it too weak to
observe.
After the preliminary study, the reaction between catechol and other anilines was
conducted. The results of these studies are summarized in Table 25. The reaction of
catechol and anilines in the presence of laccase and lipase PS provided a higher yield
than the reaction in presence of laccase only. The yield of the product in the reaction with
lipase PS increased up to 70% compare to the reaction without lipase PS (Table 25, Entry
4). Therefore, the overall yield of the product of this reaction system can be enhanced by
lipase PS.
215
Figure 113. Proposed reaction pathway of laccase/lipase catalytic system for the reaction between catechol (5a) and aniline (10).
OH
OH
5a
O
OLaccase
Lipase
10
HO
O
NH2
NH
HO
HO NH
10
NH2Lipase
NH
HO
HO N
N
Oxidation
11a, MW = 290
O2 H2O
216
Table 25. Reactions of catechol and anilines in the presence of laccase,with (or without) lipase PS in aqueous medium.
Yielda of Product (%) Entry R1
Without Lipase With Lipase
1 H 23 30 2 H 23 28b 3 OCH3 25 37 4 Cl 30 51 5 CH3 32 50 a Isolated yield; b Used 1848U of lipase PS.
OHHO
NN R1R1
HO
OH NH2
R1
Laccase, Lipase PS
Phosphate buffer pH 7rt, 3.5 h
+
5a 10 11
11a: R1 = H11b: R1 = OCH311c: R1 = Cl11d: R1 = CH3
217
7.4 Conclusions
In conclusion, this study demonstrates the potential of using lipase to catalyze
Michael addition reaction, and presents a new co-catalytic enzymatic system employing
laccase and lipase for green chemistry synthesis. Lipase was found to catalyze the
addition reaction between laccase-generated o-quinones and 1,3-dicarbonyl compounds
in aqueous medium. In this reaction, the catalytic system of laccase and lipase PS was
regioselective, providing only one isomer product and is the first example of a two
enzyme catalytic system for the synthesis of benzofurans. The yields of the products from
reaction depend on the reactivity of the starting catechols and β-dicarbonyl compounds.
Based on our experimental results, catechols with moderate reactivity yield benzofuran
products in excellent yield. Moreover, lipase was also shown to catalyze the addition
reaction between laccase-generated o-quinone and aromatic amines. In the presence of
lipase and laccase, the yield of the final products increased in the range from 30 to 70%
when compared to the reaction in the presence of laccase alone. Therefore, this paper
illustrates a unique aqueous-based two-enzyme system for green chemistry synthesis and
future applications are under study.
218
CHAPTER 8
MODIFICATION OF HIGH-LIGNIN CONTENT SOFTWOOD
KRAFT PULP WITH LACCASE AND AMINO ACIDSv
8.1 Introduction
The interest in modifying cellulosic fibers especially with the assistance of
enzymes is a growing field of research and interest [262]. A variety of enzymes are
available for the surface modification of lignocellulosics fibers [263,264]. Compared to
chemical treatments which involve harsh reaction conditions, loss of desirable
components, and potential use of hazardous chemicals, enzymatic treatment conditions
are often milder, less damaging to the fiber, and are environmentally friendly. Enzymatic
surface modifications of fibers can be accomplished with glucohydrolysis and oxidative
enzymes [263]. One of these oxidoreductases is laccase (benzenediol :oxygen
oxidoreductase, EC 1.10.3.2) which is a multi-copper-containing oxidoreductase enzyme
widely distributed in plants and fungi [3]. The majority of fungi that produce laccase
belong to the class of white rot fungi involved in lignin degradation and can mineralize
this substrate. Laccase can catalyze the oxidation of various substrates including phenols,
vThis manuscript was accepted for publication in Enzyme and Microbial Technology, 2008. It is entitled as “Modification of high-lignin content softwood kraft pulp with laccase and amino acids”. The other author is Dr. Arthur J. Ragauskas from the School of Chemistry and Biochemistry at the Georgia Institute of Technology
219
benzenediols, aminophenols, polyphenols, polyamines, and lignin-related molecules, with
concomitant reduction of oxygen to water [4-10].
Laccase applications in pulp and paper technology have been reported for
biopulping, biobleaching, deinking, mill process water and effluent treatment, and fiber
modification [20]. Recently, laccase research studies have shifted toward fiber
modification. Laccase has been used to catalyze biografting of a variety of substrates to
technical lignins. For example, Lund and Ragauskas demonstrated that laccase catalyzed
the grafting of guaiacol sulfonate to lignin which enhanced its water solubility [22].
Huttermann et al. reported that laccase can catalyze the reaction of lignin with cellulose
yielding a product in which the lignin was covalently bounded to cellulose [23].
Furthermore, Mai et al. grafted lignin with synthetic polymers derived from acrylic and
acrylamide to create a new class of engineered plastics [24-27]. In addition, laccase has
been shown to have the potential to biograft low-molecular-weight compounds to lignin-
rich cellulosic fibers. Viikari et al. [28] recently modified the fiber surfaces of
thermomechanical pulp (TMP) by laccase and tyramine. This modification is a two-stage
functionalization method consisting of enzymatic activation of fiber surfaces followed by
addition of radicalized compounds reacting preferentially by radical coupling. Chandra et
al. reported the grafting of phenolic acids, including 4-hydroxyphenylacetic acid (PAA)
[30], 4-hydroxybenzoic acid (4-HBA) [31], and gallic acid [29] to high-lignin content
softwood kraft fibers. The grafting of these charged phenolics via a laccase generated
phenolate radical was shown to lead to improved tensile and burst strength for the
resulting paper. The paper strength improvements were ascribed to the capacity of
carboxyl groups to promote fiber-fiber bonding and fiber swelling [406-412].
220
Laccase is also attractive for fine chemical synthesis because of its high stability,
selectivity for phenolic substructures, and mild reaction conditions [11-
14,18,19,244,366,379,380,404,413]. For instance, Michałek and Szarkowska [413]
studied the reaction between laccase generated p-quinones and amino acids to produce
quinone-amino acid complexes. The propensity of laccase to catalyze the oxidation
polyphenolic has been reported by Chakar and Ragauskas [366] and Lalk et al. [11] has
reported a laccase catalyzed nuclear animation reaction with p-diphenols and aromatic
amines. According to the studies in Chapter 4-7, laccase has also been shown to initiate a
cascade synthesis of naphthoquinone derivatives via Diels-Alder reaction between
benzenediols and dienes [379,380] as well as the synthesis of benzofuran derivatives via
oxidation-Michael addition between o-benzenediols and 1,3-dicarbonyl compounds
[404]. Based on these results, it was apparent that laccase can be employed to generate
reactive quinoidal structures in lignin-rich fibers that could then be reacted with amino
acids to generate enhanced fiber charge as shown in Figure 114. This Chapter examines
the optimal grafting conditions with respect to fiber charge and its impact on sheet
strength properties.
221
Figure 114. Propose mechanism for the grafting treatment of high-lignin content softwood kraft pulp with laccase and amino acids.
8.2 Experimental Section
8.2.1 Materials
All chemicals were obtained from Aldrich and used as received without further
purification. Laccase (EC 1.10.3.2) from Trametes villosa was donated by Novo Nordisk
Biochem, North Carolina and frozen till used. A commercial linerboard softwood kraft
pulp (17% of lignin content) was obtained from a southeastern U.S.A manufacturing
facility. The pulp was exhaustively washed until the filtrate was pH neutral and
colorless. Pulp was air dried and soxhlet extracted for 24 hours with acetone with
subsequent washing with water prior to all treatments.
Laccase (red)
Laccase (ox)
OH
OH
Lignin
OH
O
Lignin
O
O
Lignin
O2
H2O
OH
OH
Lignin
H2N COOH
R
Addition ReactionHNHOOC
R
222
8.2.2 Enzyme Assay
Enzyme activity measurement is described in Chapter 3 (Experimental Materials
and Procedures).
8.2.3 Pulp Treatment
Laccase (80 U/1 o.d. g pulp) and an amino acid (3.2 mmol/1 o.d. g pulp) were
added with stirring to a 5% consistency aqueous suspension of linerboard pulp buffered
to pH 7 with 0.10 M sodium phosphate solution. The resulting slurry was stirred for 4 h
at room temperature and then left stand 20 h. After treatment, the pulp sample was
filtered, washed with deionized water until the filtrate was colorless and air-dried.
Typically, pulp mass recovery was 95%.
8.2.4 Bulk Acid Group Measurment
Conductrometric titration for bulk acids was based on the work of Katz [328]. In
brief, pulp (1.50 g o.d.) was stirred in 300.00 ml of 0.10 M HCl for 1 hour followed by
rinsing in a fine fritted funnel with deionized water. The sample was then re-suspended in
250.00 ml of 1 mM NaCl solution, spiked with 1.50 ml of 0.10 M HCl and titrated
against 0.05 M NaOH at 0.25 ml increments, recording the conductivity at each
increment. The titration data was plotted as conductivity vs. volume to determine the
milli-equivalent of acid groups per g of pulp. The reported results were the average of
two measurements which typically differed by less than 3%.
223
8.2.5 Paper Testing
Treated pulps and control were disintegrated for 30,000 revolutions and then were
refined for 3,000 revolutions according to TAPPI Standard T 248 [327]. Handsheets (3 g)
were formed according to TAPPI Standard T 205 [327] and TAPPI conditioned (23 ˚C,
50% relative humidity) for at least 24 hours before physical testing.
Apparent density, tensile strength, tearing resistance, and wet tensile strength
were determine according to TAPPI methods T 210, T 494, T 414, and T 456 [327]. The
results of each physical testing were the average of five measurements with error less
than 3%. Nitrogen content was analyzed by elemental microanalysis (Huffman
Laboratories, Inc., Golden, CO) and the results are reported on a dried sample basis. The
SEM pictures of handsheets were taken using a Hitachi S-800 FE-SEM. The handsheet
sample was stuck on the SEM sample holding stub by the conductive double sides sticky
carbon film and then was coated with alloy of Au/Pt prior to analysis.
8.3 Results and Discussion
8.3.1 Preliminary Study of the Grafting Condition
To determine the optimal condition for the modification of the linerboard pulp, a
preliminary study was conducted with laccase and glycine (Gly). In this modification, the
linerboard pulp was first stirred at 5% consistency in a pH 7.0 phosphate buffer solution
with laccase (80 U/1g pulp) and Gly (0.8 mmol/1g pulp) for 4 h at room temperature and
224
then left unstirred for an additional 20 h. The treated pulp was washed, filtered, air dried,
and then analyzed for bulk fiber charge. The results of analysis for laccase-Gly treated
pulp (Lac/Gly), laccase-treated pulp (Lac) Gly-treated pulp (Gly) and control pulp are
shown in Fig. 2a. These results demonstrate that laccase treated pulp provided a higher
yield of acid groups compared to the control pulp due to the oxidation of lignin by
laccase. Gly-treated pulp gave the similar acid content compared to the control pulp. This
result suggested that Gly itself did not react with the lignin in the pulp fibers under the
reaction conditions employed. However, when the pulp was treated with both laccase and
Gly, the treated pulp gave the highest yield of carboxyl groups. This increase of carboxyl
groups indicated that laccase-treated fibers facilitated the grafting of Gly onto the fiber
lignin. Then, to determine the effect of the treatment conditions on grafting, the pH of the
treatment was changed from 7.0 to 4.5 which is known to be the optimal pH for laccase
[365,366]. The result shows that the treatment at pH 4.5 provided a reduced content of
acid groups than the treatment at pH 7.0. This was attributed to the higher pH
requirements needed for Micheal addition of amino acids to lignin quinonoid compounds
(Figure 115 (top)). The requirement of using higher pH, pH 7, for the Micheal addition
catalyzed by laccase was also reported by Michałek et. al. [413] and Ragauskas et al.
[404].
225
0.1
0.11
0.12
0.13
0.14
0.15
0.16
0.17
0.18
0.19
0.2
ControlPulp
Lac Gly Lac/GlypH7.0, RT
Lac/GlypH4.5, RT
Lac/GlypH7.0, 45 ºC
Bulk Acid Group
s (m
eq/g)
0.17
0.175
0.18
0.185
0.19
0.195
0.2
0.8 mmol 1.6 mmol 2.4 mmol 3.2 mmol
Bulk Acid Group
s (m
eq/g)
1st Measurement 2nd Measurement
Figure 115. (top) Bulk acid group content of control pulp, laccase treated pulp (Lac), glycine treated pulp (Gly), and laccase-glycine treated pulp (Lac/Gly) at different conditions (The control pulp, laccase treated pulp and Gly-treated pulp were treated in the same condition as laccase-Gly treated pulp but no laccase and Gly, no Gly, and no laccase, respectively); (bottom) bulk acid group content of pulps treated with laccase and different amount of glycine at pH 7.0 and room temperature.
226
The effect of the reaction temperature on this grafting procedure was also
examined. The pulp was treated at pH 7.0 and at 45 ˚C instead of at room temperature.
The result of this treatment showed that the increase in temperature did not increase the
acid group content of the fibers (Figure 115(top)). Therefore, the optimal condition of
this fiber modification was the treatment at pH 7.0 and at room temperature. The effects
of differing charges of Gly were also evaluated as shown in Figure 115(bottom). These
results shows that the pulp treated with 1.6 mmol of Gly showed similar amount of bulk
acid content when compare with 0.8 mmol Gly-treated pulp. However, the acid content
increased when the pulp was treated with 2.4 mmol and 3.2 mmol of Gly/1 g fiber.
8.3.2 The Effect of Amino Acids on the Modifying Fibers
After this preliminary study, the next phase was to examine the effect of differing
amino acids for laccase initiated fiber grafting. Softwood linerboard kraft pulp was
treated with laccase (80 U/1g pulp) and amino acid in phosphate buffer pH 7.0 at room
temperature. A variety of amino acids were used for this study including Gly,
phenylalanine (Phe), serine (Ser), aspartic acid (Asp), histidine (His), arginine (Arg), and
alanine (Ala). The properties of amino acids mainly depend on the pH of the surrounding
environment. The amino acids can become more positively or negatively charged due to
the loss and gain of protons (H+) at a given pH. In general, the pK values of the α-
carboxylic acid groups of amino acids lie in a small range around 2.2 so that above pH
3.5 these groups are almost entirely in their carboxylate forms. The α-amino groups all
have pK values near 9.4 and are therefore almost entirely in their ammonium ion forms
below pH 8.0 [414]. Therfore, at the experimental pH (pH 7.0), both the carboxylic acid
227
and the amino groups of α-amino acids are ionized. When the amino acids have charged
polar side chains, the pK values of the side chain groups have to be considered. In this
study, histidine side chain, an imidazolium moiety (pK = 6.0), was deprotonated at pH
7.0. Therfore, the histidine side chain can participate in the addition reaction with the
laccase-oxidized fibers at this pH. The results illustrated in Figure 116a show that His
gave the highest acid content compared to the other amino acids. This result was ascribed
to the enhanced nucleophilic property of the nitrogen of imidazole side chain of His.
Moreover, when considered the isoelectric point (pI) of the amino acids with nonpolar or
uncharged side chains, including Gly (pI = 6.06), Ala (pI = 6.01), Phe (pI = 5.49), and
Ser (pI = 5.68), their pI are all below 7. Therefore, at the pH above their pI (pH 7.0),
some of ammonium ions of these amino acids were deprotonated which led to the
liberation of some free amino groups that can react with the oxidized fibers. As a
consequence, the acid groups of the fibers increased in some content after the treatment
with these amino acids and laccase at pH 7.0 (Figure 116a).
In addition, different amounts of each amino acid (i.e., 1.6, 2.4, and 3.2 mmol/1g
pulp) were examined to find the optimal amount of amino acid for modifying fibers. The
results in Figure 116a also indicate that the greater the amount of amino acid employed
the greater increase in fiber charge for most amino acids. The acid group content reached
the maximum yield when the amount of amino acids was 3.2 mmol/1g pulp. Therefore,
3.2 mmol/1g pulp was chosen as an optimal amount of amino acids for this treatment
system.
Next, the interaction between amino acids and pulp fibers was investigated. The
pulp was treated with an amino acid (3.2 mmol/1g pulp) only and compared the acid
228
content with control pulp, laccase-treated pulp and laccase-amino acid treated pulp.
Figure 116b demonstrates that the amino acid-treated pulp provided a 10-25% increase of
carboxyl group content compared to control pulp. These results indicate that some of
amino acid can react with pulp fibers presumably due to quinonoid structures present in
kraft pulps [415]. However, the carboxyl group content of the amino acid treatments was
still 11-20% less than of the laccase-amino acid treatments. Therefore, the highest acid
group content was obtained when the linerboard pulp was treated with both laccase and
amino acid.
229
0.15
0.16
0.17
0.18
0.19
0.2
0.21
0.22
Bulk Acid Gro
ups (m
eq/g
)
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
1.6 mmol
2.4 mmol
3.2 mmol
Gly Phe Ser Asp His Arg Ala
Amount of amino acid (mmol/g pulp)
1st Measurement 2nd Measurement
0.13
0.14
0.15
0.16
0.17
0.18
0.19
0.2
0.21
0.22
Bulk Acid Gro
ups (m
eq/g
)
Control Pulp La
cGly
Lac/G
ly Phe
Lac/P
he Ser
Lac/Ser As
p
Lac/A
sp His
Lac/H
is Arg
Lac/A
rg Ala
Lac/A
la
1st Measurement 2nd Measurement
Figure 116. Bulk acid group content of (a) linerboard pulps treated with a variety of amino acids in the presence of laccase (80 U/1g pulp); (b) linerboard pulps treated with a variety of amino acids (3.2 mmol/ 1g pulp) in the presence and absence of laccase.
(a)
(b)
230
8.3.3 The Effect of Laccase Dose
After optimizing the treatment condition, the next study was to determine the
effect of laccase dose on the modifying fibers. The experiments were conducted by
treating linerboard pulp with different amount of laccase which are 20, 40, 60, 80, and
100U/1g pulp in the presence of His (3.2 mmol/1g pulp) in phosphate buffer pH 7.0 at
room temperature. Figure 117 demonstrates that the carboxyl group content increased
when the amount of laccase increased. The carboxyl group content reached the highest
amount when the amount of laccase was 80 U/g pulp. Therefore, the optimal amount of
laccase for this modification was 80 U/g pulp.
Effect of Laccase Dose
0.17
0.175
0.18
0.185
0.19
0.195
0.2
0.205
0.21
0.215
0.22
20 U 40 U 60 U 80 U 100 U
Activity of Laccase/1g pulp
Bulk Acid Group
s (m
eq/g)
1st Measurement 2nd Measurement
Figure 117. Bulk acid group content of linerboard pulps that were treated with histidine (3.2 mmol/ 1g pulp) and different amount of laccase.
231
8.3.4 Nitrogen Content of Laccase-His Treated Pulp
The laccase-His grafting treatment conditions which provided the best yield of
bulk fiber acid groups were selected for further study. The linerboard pulp was treated
with laccase and histidine using the optimal condition as described in experimental
section 8.2.3. Then, the pulp samples were sent for nitrogen analysis. Nitrogen content of
laccase-His treated pulp was measured and compare with nitrogen content of control and
laccase treated pulp. The nitrogen content of laccase-His treated pulp was 120-140%
higher than of control and laccase treated pulp as shown in Figure 118. These results
show that His was bonded with pulp fibers after the grafting treatment which led to the
increase of nitrogen content of the fibers.
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
Control pulp Lac Lac/His
%Nitrogen
Figure 118. Nitrogen content of control pulp, lacccase treated pulp (Lac), and laccase-His treated pulp (Lac/His).
232
8.3.5 Paper Strength Properties
The objective of this section is to evaluate the effects of the laccase-amino acid
grafting treatment on paper strength properties. The physical properties of handsheets
made from laccase-His treated pulp were compared to the physical properties of the
handsheets made from control pulp and laccase treated pulp. The results of the paper
testing are illustrated in Figure 119. The strength properties of the handsheets were
examined including tensile strength, tearing resistance, and wet tensile strength. These
results indicate that the handsheets made from laccase-His treated pulp gave the highest
strength properties in comparison to handsheets made from control and laccase treated
pulp. The ratio of wet/dry strength is about 5.2 for the laccase-His treated pulp. Although
it has been suggest that the minimum ratio of wet/dry strength about 15 is required for the
wet-strength paper [416], this study is a good start for the modification of lignocellulosic
fibers by laccase via oxidation-Michael addition. Therefore, in the future, the further
investigation to improve the wet tensile strength of resulting paper for this modification
system has to be conducted. The improvement of wet tensile strength of unbleached kraft
pulp by the combination of lacccase with mediator and a heat treatment has been reported
by Lund and Felby [297]. The wet/dry strength ratio of laccase, laccase-mediator, and
laccase-mediator with heat treatment is 3.5, 6.7, and 14.7, respectively. This shows that
heat treatment has a tremendous effect on the increase of wet strength property.
Compared to Lund and Felby’s study, our wet/dry strength ratio is comparable to those
results without heat treatment. Therefore, our fiber modification system could be further
improved by using laccase in combination of mediator or heat treatment to increase the
wet tensile strength of the modified fibers.
233
Moreover, the images of the handsheet surface of the control, laccase treated, and
laccase-His treated pulp were taken by the scanning electron microscope (SEM). SEM
images in Figure 120 show that the laccase-His treated fibers are more collapse than
control and laccase treated fibers which led to form better bonding between fibers in
handsheet resulting in the increase of the paper strength of laccase-His treated pulp.
234
(a) Tensile Strength
50
51
52
53
54
55
56
57
Control Pulp Lac Lac/His
Tensile
Inde
x (N.m
/g)
(b) Tear Strength
12
12.5
13
13.5
14
14.5
15
15.5
16
Control Pulp Lac Lac/His
Tear In
dex (m
N.m
2 /g)
(c) Wet Tensile Strength
1
1.5
2
2.5
3
3.5
Control Pulp Lac Lac/His
Tensile
Inde
x (N.m
/g)
Figure 119. Physical paper properties of handsheets made from control pulp, laccase treated pulp (Lac), and laccase-histidine treated pulp (Lac/His); (a) tensile strength; (b) tear strength; (c) wet tensile strength.
235
Figure 120. Scanning electron microscope (SEM) images of handsheets made from (a) control pulp; (b) laccase treated pulp; (c) laccase-histidine treated pulp.
236
8.4 Conclusions
This study presents a new environmentally-friendly method for modifying lignin-
rich fibers. This modification employed laccase to oxidize lignin in the fibers, and then
the carboxyl groups were introduced to pulp fibers by an addition reaction between the
oxidized fibers and amino acids. The condition for this treatment was pH 7.0 at room
temperature. Laccase-amino acid treatment of fibers resulted in an increase in carboxyl
group content of the fibers that enhanced the strength properties of the resulting paper,
including tensile strength, tearing resistance, and wet tensile strength. The SEM images
show that the laccase-amino acid treated fibers are more collapse than control and
laccase-treated fibers which led to form better bonding between fibers in handsheet. In
this study, among the several different amino acids studied, the treatment of pulp with
laccase and His provided the best result in increasing carboxyl group content and paper
properties. The ability to use laccase selectively graft amino acids to lignin rich pulp
fibers provides a new and unique fiber modification technology which will have many
future opportunities. The improvement of this fiber modification system to increase the
strength properties of the modified paper is under investigated.
237
CHAPTER 9
OVERALL CONCLUSIONS
The original idea about using laccase for this study was inspired by various
interesting applications of laccase as biocatalysts. Laccase has been known to have
applications in many industrial areas, expecially in the pulp and paper industry.
However, the applications of laccase have recently shifted toward fine chemical
synthesis because of its high stability, selectivity for phenolic substructures, and mild
reaction conditions. This study utilized the oxidative potential of laccase to convert
hydroquinones to quinones in situ. Since the quinonoid compounds have a wide
spectrum of chemistry, various possible reactions of the in situ-generated quinones
can be investigated. First, the property of quinoniod compounds as good dienophiles
for the Diels-Alder reactions attracted our interest. Moerover, many studies showed
that the Diels-Alder reactions performed in an aqueous medium showed beneficial
effects on the reaction rate, reactivity, and selectivity of Diels-Alder reaction.
Therefore, the study of the laccase-triggered Diels-Alder reaction in aqueous media
was conducted first. This reaction methodology provides a unique green chemistry
synthesis.
In Chapter 4, the para-quinones were generated in situ by the laccase oxidation
of the corresponding 1,4-hydroquinones and subsequently underwent the Diels-Alder
reaction with dienes, and further oxidation to finally generate 1,4-naphthoquinones, in
good yields. However, the reactivity of the reaction depends on the substrate specificity
of laccase and the reactivity of both generated quinones and dienes. Temperature also has
238
an important impact on the formation of the final products. To obtain the
naphthoquinones as major products, the reactions have to perform at 70 oC. At the lower
temperature, 25 oC, the major products showed to be the Diels-Alder adducts. This
successful synthesis of p-naphthoquinones catalyzed by laccase led to the further study of
the laccase-triggered Diels-Alder reaction for o-naphthoquinones synthesis in Chapter 5.
This study has to deal with the very reactive in situ-generated o-quinones that easily
undergo dimerization and polymerization. Therefore, the reactions were conducted at a
low temperature (3-25 oC) to lower the rate of those side reactions and a high excess of
dienes were used to push the reaction toward Diels-Alder reaction. In addition, these
reactions were carried out in an aqueous medium and yielded o-naphthoquinones up to
80%, depending on the exact structure of the starting hydroquinone and diene.
Besides Diels-Alder reactions, Michael addition reactions of in situ-generated o-
quinones were also investigated. In Chapter 6, the cascade synthesis of benzofuran
derivatives was conducted from the reaction of catechols and 1,3-dicarbonyl compounds
via oxidation-Michael addition in the presence of laccase and Sc(OTf)3/SDS. In this
procedure, ortho-quinones, generated in situ from the oxidation of catechols by laccase,
underwent the Michael addition reaction with 1,3-dicarbonyl compounds, and then
underwent intramolecular cyclization to benzofuran derivatives. This reaction was carried
out under air at room temperature, in an aqueous medium, and provided benzofuran
products in 50 – 79% yield. In addition, this reaction system showed recyclability.
Although the use of an aqueous Lewis acid was critical for efficient synthesis of the
desired compounds, this system still produced a hazardous waste from the transitional
metal catalyst. Therefore, to enhance the overall green chemistry aspect, the use of lipase
239
as an alternative catalyst in conjunction with laccase as an alternative methodology for
the synthesis of benzofuran derivatives was developed in Chapter 7. This laccase/lipase
reaction system was carried out under air at room temperature, in an aqueous medium,
and provided benzofuran products in good yields. Moreover, this laccase/lipase co-
catalytic system was also used to catalyze the Michael addition of in situ-generated o-
quinones and anilines. In the presence of lipase and laccase, the yield of the final
products increased in the range from 30 to 70% when compare to the reaction in the
presence of laccase alone. Therefore, this study illustrates a unique aqueous-based two-
enzyme system for green chemistry synthesis.
In the last phase of this research, the interest shifted toward another interesting
application of laccase, which is fiber modification. Laccase has been reported to facilitate
the grafting of a variety of compounds to lignin or lignocellulosic fibers. Chapter 8
demonstrates the potential of laccase-facilitated grafting of amino acids to high lignin
content pulps to improve their physical properties in paper products. These physical
properties can be enhanced by increasing ionic fiber charges. In an effort to increase
carboxylic acid groups, a unique two-stage laccase grafting protocol, in which fibers were
initially treated with laccase followed by grafting reactions with amino acids was
developed. The condition for this treatment was pH 7.0 at room temperature. In this
study, a variety of amino acids, including glycine, phenylalanine, serine, arginine,
histidine, alanine, and aspartic acid, were examined. The results show that histidine
provided the best yield of acid groups on pulp fiber and was used for the preparation of
handsheets for physical strength testing, including tensile, tear, and wet tensile strength
properties. Laccase-histidine treated pulp showed an increase in strength properties of the
240
resulting paper. Moreover, this study presents a new environmentally-friendly method for
modifying lignin-rich fibers.
241
CHAPTER 10
RECOMMENDATIONS FOR FUTURE WORK
Several other studies might be conducted to further explore other applications of
laccase, both in organic synthesis and in fiber modification. Some particularly attractive
options are as follows:
To address the environmental concern, immobilized laccase would be used
in the reaction. The immobilized laccase could be reused and would
reduce waste from the reaction.
The use of laccase alone in the reaction limits the scope of substrates. The
addition of laccase mediators, such as ABTS, HBT, and TEMPO, into the
reaction system would broaden the scope of the substrates and would lead
to the discovery of new green synthetic chemistry.
According to this research, reaction conditions, such as temperature and
pH, affect the formation of the reaction products. Therefore, conducting
the reaction at different conditions could provide different final products
and could lead to the discovery of new compounds.
Laccase could be used to facilitate the grafting of other compounds to
high-lignin content pulp fibers to improve the properties of existing
products and create new product platforms.
Future research programs should focus on large-scale laccase-biografting
technology.
242
APPENDIX A
NMR AND IR SPECTRA OF NEW COMPOUNDS
A.1 NMR and IR Spectra of New Compounds in Chapter 4
There are two new compounds obtained from the experiments in Chapter 4:
1,4-Dihydro-6-methoxy-1,4-ethanonaphthalene-5,8-dione (3f)
1,4-Dihydro-6-bromo-1,4-ethanonaphthalene-5,8-dione (3h)
O
O
MeO
O
O
Br
243
A.1.1 1H-NMR Spectrum of compound 3f
244
A.1.2 13C-NMR Spectrum of Compound 3f
245
A.1.3 IR Spectrum of Compound 3f
246
A.1.4 1H-NMR Spectrum of Compound 3h
247
A.1.5 13C-NMR Spectrum of Compound 3h
248
A.1.6 IR Spectrum of Compound 3h
249
A.2 Spectra of New Compounds in Chapter 5
There are two new compounds obtained from the experiments in Chapter 5:
4,7,8-trimethyl-1,2-naphthoquinone (6e)
4-methyl-6,7-dimethoxy-1,2-naphthoquinone (6f)
O
CH3
O
CH3
CH3
O
CH3
O OMe
OMe
250
A.2.1 1H-NMR Spectrum of Compound 6e
251
A.2.2 13C-NMR Spectrum of Compound 6e
252
A.2.3 1H-NMR Spectrum of Compound 6f
253
A.2.4 13C-NMR Spectrum of Compound 6f
254
A.3 Spectra of New Compounds in Chapter 6
There is one new compound obtained from the experiments in Chapter 6:
Ethyl-5,6-dihydroxy-2,7-dimethyl-3-benzofuran carboxylate (9d).
O
Me
OEtO
Me
OH
OH
255
A.3.1 1H-NMR Spectrum of Compound 9d
256
A.3.2 13C-NMR Spectrum of Compound 9d
257
A.3.3 HSQC Spectrum of Coumpound 9d
258
A.3.4 HMBC Spectrum of Compound 9d
259
A.4 Spectra of New Compounds in Chapter 7
There are three new compounds obtained from the experiments in Chapter 7:
Compound 11b
Compound 11c
Compound 11d
N N
HO OH
MeO OMe
N N
HO OH
Cl Cl
N N
HO OH
H3C CH3
260
A.4.1 1H-NMR Spectrum of Compound 11b
261
A.4.2. 13C-NMR Spectrum of Compound 11b
262
A.4.3 HMQC Spectrum of Compound 11b
263
A.4.4 HMBC Spectrum of Compound 11b
264
A.4.5 1H-NMR Spectrum of Compound 11c
265
A.4.6 13C-NMR Spectrum of Compound 11c
266
A.4.7 HMQC Spectrum of Compound 11c
267
A.4.8 HMBC Spectrum of Compound 11c
268
A.4.9 1H-NMR Spectrum of Compound 11d
269
A.4.10 13C-NMR Spectrum of Compound 11d
270
A.4.11 HMQC Spectrum of Compound 11d
271
A.4.12 HMBC Spectrum of Compound 11d
272
A.4.13 IR Spectra of compound 11b, 11c and 11d
Compound 11b Compound 11c Compound 11d
273
APPENDIX B
COPYRIGHT PERMISSION
B.1 Permission of RSC (Green Chemistry)
274
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APPENDIX C
TENSILE AND TEAR STRENGTH
C.1 Tensile Strength
The tensile strength of paper sheets is especially complex as many variables play
a role in controlling the magnitude of this property. Tensile strength is dependent on both
the fiber strength properties and the bonding that occurs between fibers. The tensile
strength theory that has attracted the most attention has been that of Page. The “Page”
equation (Equation 4) was shown in a publication in 1969 [417] and remains a fixture in
paper physics discussions. The equation represents a comprehensive account of the
variables encountered in attempting to predict tensile strength from the properties of the
fiber and for bonds between fibers. The equation (Equation 4) also attempts to calculate
“bondstrength” from all of these variables affecting tensile strength
(1/T) = (9/8Z) + [(12g × C)/(P × l × b × RBA)] Where: l = fiber length (length) b = fiber-fiber bond strength (N/m2) RBA = relative bonded area (unit less) g = gravitational constant -(length/second2 = 9.8 m/s2) T = tensile breaking length (length) Z = zero span tensile (length) C = fiber coarseness (weight/length) P = fiber perimeter (length)
Equation 4. The Page equation.
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Equation 4 shows that the inverse of tensile strength should be linearly
proportional to the inverse of fiber strength, fiber length, and RBA. The tensile strength
predictions of the Page equation are illustrated in Figure 121.
Figure 121. Predictions from Page equation for tensile strength of paper vs. relative bonded area together with the qualitative effect of increasing fiber properties.
The relative bonded area (RBA) in Page’s equation is a measure of the contact
area between fibers in the sheet [417].This is measured by light scattering co-efficient or
through nitrogen absorption measurements. Increases in bonded area can be achieved by
increasing wet-pressing pressure. With subsequent testing of a strength property (such as
tensile strength) and scattering coefficient, the sheet strength can be extrapolated to zero
sheet strength. The result of this extrapolation is an estimate of the scattering coefficient
of unbonded fibers (So) that can be used to calculate the relative bonded area [417].
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Equation 5 shows the relationship between relative bonded area and light scattering
coefficient.
RBA = (So-S)/So Where: So = scattering co-efficient of the unbonded sheet (m2/kg) S = scattering co-efficient for a paper sheet (m2/kg) Equation 5. Page’s equation for computing relative bonded area.
In the Page equation, most of the variables are measurable except for b, the fiber
to fiber bond strength or “shear strength” of the fiber bonds [417]. Once all of the
measurable variables are obtained, the Page parameter ([1/T – 9/(8Z)]-1) can be plotted
against the light scattering coefficient (S) (Equation 6). This plot can be used to obtain
the bond strength (b) and the scattering coefficient of the unbound fibers (So) from the
slope and intercept respectively.
[(1/T) – (9/8Z)]-1 = b × [(1/ γ) – (S/( γ × So))] γ = [(12g × C)/ (P × l)] Equation 6. Parameters to plot for obtaining bond strength using the Page equation.
The Page equation is only valid for sheets made with good formation, free from
kinks or curls [418]. This is because sheets with poor formation fail earlier due to uneven
concentrations of stress in areas of low basis weight. Kinks and curls cause changes in the
283
fiber length variable in the equation. The kinks and curls also decrease the number of
load-bearing elements in the sheet.
In the physical testing of paper, tensile strength is determined by measuring the
force required to break a narrow strip of paper where both the length of the strip and the
rate of loading are closely specified [285]. The amount of stretch at rupture may be
determined at the same time. Some modern testers provide a plot of the stress/strain curve
and compute the area under the curve which is referred to as tensile energy absorption, a
measure of paper toughness. These testers also provide for measurement of creep under
various tensile loading.
C.2 Wet Tensile Strength
Paper is a layered mat consisting of a network of cellulose fibers held together by
intermolecular forces (van der Waals and hydrogen bonding) which are very sensitive to
water. The extent of bonding steadily decreases as the water content of the paper
increases. The water wets the fibers, and then, the bonds are broken leaving somewhere
between 3% and 10% of the original dry strength (at 50% relative humidity). The residual
strength of wet paper results from remaining covalent fiber-fiber bonds. Therefore, there
is a need for paper products to retain some strength when subjected to high humidity or
when soaked in water. Many applications have been developed to improve the wet
strength of paper [416].
The way to determine wet strength of the paper is to measure its burst or tensile
strength when wet. There are useful Standard Methods for the determination of wet
284
strength (e.g. TAPPI Method T456), although many non-standard tests have been
developed over the years. In the TAPPI Method, a strip of paper is completely wetted
before applying a breaking force. The paper is immersed in water or, if it is too weak, it is
mounted in the jaws of a tensile tester and wet midway over a distance of 2.54 cm. The
load required to break the paper is then recorded. The result reported as percent wet
strength (wet strength as a percentage of the dry strength).
C.3 Tear Strength
Tearing resistance is the total energy per tear length consumed when a specimen
of a given geometry undergoes tearing. Tearing resistance therefore has the units of load
and is sometimes called tear strength, although it is energy, not stress, that one measures.
Tearing strength is normally determined with the Elmendorf apparatus which uses a
falling pendulum to continue a tear in the paper sample when the force is applied
perpendicular to the plane of the sheet; the loss of energy, measured by the height of
swing of the pendulum, is related to the force required to continue the tear [285]. The
Elmendorf tear test is recognized as a good measure of fiber strength within the sheet.
Apparatus for carrying out in-plane tear testing is available, but the procedure is not
widely utilized. In the in-plane tear measurement, load is applied in the plane of paper,
often at a 2 x 6° angle as Figure 122(a) shows. In the out-of-plane tear test or Elmendorf
tear of Figure 122(b), load is in the out-of-plane direction.
285
Figure 122. (a) The in-plane tear test; (b) the out-of-plane or Elmendorf tear test.
286
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